Amino-functionalized 5' cap analogs as tools for site-specific

Subscriber access provided by CORNELL UNIVERSITY LIBRARY

Article

Amino-functionalized 5’ cap analogs as tools for sitespecific sequence-independent labeling of messenger RNA Marcin Warminski, Pawel J Sikorski, Zofia Warminska, Maciej Lukaszewicz, Anna Kropiwnicka, Joanna Zuberek, Edward Darzynkiewicz, Joanna Kowalska, and Jacek Jemielity Bioconjugate Chem., Just Accepted Manuscript • Publication Date (Web): 14 Jun 2017 Downloaded from http://pubs.acs.org on June 14, 2017

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a free service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are accessible to all readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

Bioconjugate Chemistry is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Bioconjugate Chemistry

Amino-functionalized 5′ cap analogs as tools for site-specific sequenceindependent labeling of messenger RNA Marcin Warminski§,ǁ, Pawel J. Sikorski†,ǁ, Zofia Warminska†,‡,ǁ, Maciej Lukaszewicz§, Anna Kropiwnicka§, Joanna Zuberek§, Edward Darzynkiewicz§,†, Joanna Kowalska§,* and Jacek Jemielity†,* §

Division of Biophysics, Institute of Experimental Physics, Faculty of Physics, University of Warsaw, Warsaw, Poland † Centre of New Technologies, University of Warsaw, Warsaw, Poland ‡ College of Interfaculty Individual Studies of Mathematics and Natural Sciences, University of Warsaw, Warsaw, Poland * Corresponding Author Prof. Jacek Jemielity, [email protected], Centre of New Technologies, University of Warsaw, Banacha 2C, 02-097 Warsaw. Phone: +48 22 5543774, Fax: +48 22 5540801. Dr. Joanna Kowalska, [email protected], Division of Biophysics, Institute of Experimental Physics, Faculty of Physics, University of Warsaw, Pasteura 5, 02-093Warsaw. Phone: +48 22 5532342 ǁ

These authors contributed equally to this work.

ABSTRACT mRNA is a template for protein biosynthesis, and consequently mRNA transport, translation, and turnover are key elements in the overall regulation of gene expression. Along with growing interest in the mechanisms regulating mRNA decay and localization, there is an increasing need for tools enabling convenient fluorescent labeling or affinity tagging of mRNA. We report new mRNA 5′ cap analog-based tools that enable site-specific labeling of RNA within the cap using N-hydroxysuccinimide (NHS) chemistry. We explored two complementary methods: a co-transcriptional labeling method, in which the label is first attached to a cap analog and then incorporated into RNA by transcription in vitro, and a post-transcriptional labeling method, in which an amino-functionalized cap analog is incorporated into RNA followed by chemical labeling of the resulting transcript. After testing the biochemical properties of RNAs carrying the novel modified cap structures, we demonstrated the utility of fluorescently labeled RNAs in decapping assays, RNA decay assays, and RNA visualization in cells. Finally, we also demonstrated that mRNAs labeled by the reported method are translationally active. We envisage that the novel analogs will provide an alternative to radiolabeling of mRNA caps for in vitro studies and open possibilities for new applications related to the study of mRNA fates in vivo. INTRODUCTION

1 ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 2 of 35

Messenger RNA (mRNA) is a crucial mediator in gene expression, conveying the genetic information encoded in DNA to the ribosome complex, which translates the mRNA sequence to specific amino acid sequences forming proteins. mRNA is a relatively short-lived molecule, and thereby difficult to study. In bacteria, its halflife ranges from seconds to 20 minutes,(1) whereas eukaryotes have evolved an extensive mRNA processing pathway, which includes addition of a cap structure at the 5′ end and a poly(A) tail at the 3′ end (Figure 1), resulting in extension of mRNA half-life to minutes or even days.(2) Nevertheless, the concentration of mRNA in cells is usually at the nanomolar or even picomolar level,(3) making it difficult to detect and quantify mRNAs, especially in real-time experiments. To address this problem, several protocols for RNA fluorescent and affinity labeling have been developed, most of them combining chemical synthesis and enzymatic reactions.(4, 5) As RNA is a linear polymer, the main challenge in RNA labeling is to attach a tag in a single specific region so that the resulting conjugate retains (most of the) biological functions of transcript. The RNA conjugates obtained by random labeling protocols, such as commercially available labeling kits, can be successfully used for in vitro visualization and tracking, but they are usually less useful in structural and mechanistic studies, such as mRNA maturation, translation and degradation pathways. Randomly labeled conjugates can also be obtained by in vitro transcription in the presence of modified nucleoside triphosphates (NTPs) in addition to the natural NTPs.(6-11) An interesting variation to this approach, addressing the site-specificity issue, is application of unnatural base pairs displaying intrinsic fluorescence, although this requires an appropriately modified DNA template.(12-17) Other site-specific RNA labeling approaches include use of hybridization probes(18, 19) and aptamers,(20, 21) ribozyme-based covalent labeling,(22, 23) DNA-templated function transfer,(24) and post-transcriptional enzymatic modifications.(25-27)

Figure 1. Structure of eukaryotic mRNA 5′ end (m7G cap). An interesting target for site-specific mRNA labeling is the 5′ cap. The 7-methylguanosine (m7G) cap is a structural feature characteristic of all eukaryotic mRNAs, distinguishing them from other types of RNA. The cap participates in many important steps in mRNA processing and turnover, including splicing, nuclear export, initiation of translation, and decay. As such, RNA cap labeling appears to be an attractive approach for creating tools that could facilitate studies into such cap-related processes. Several approaches have already been developed to label mRNA within the 5′ cap, and such labeled molecules have been employed in in vitro and in vivo experiments. For example, it has been shown that several site-specific N7- or N2- guanine methyltransferases engineered to accept synthetic analogs of S-adenosylmethionine as cofactors could transfer reactive functional


Page 3 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


groups onto the RNA cap, which could be subsequently labeled using, for example, ‘click’ chemistry reactions and visualized by fluorescence methods.(28-32) However, the introduction of bulky fluorescent dyes at the methyltransferase labeling sites inhibited mRNA translation.(33) Another example of enzymatic labeling is the use of fluorescently labeled GTP derivatives in RNA-capping reaction catalyzed by Vaccinia Capping Enzyme (VCE); this approach is limited by specificity of the capping enzymes used. Finally, RNA labeling can be achieved by introduction of properly designed, labeled dinucleotide cap analogs at the RNA 5′ end by standard in vitro transcription protocols. Examples include incorporation of cap analogs labeled with biotin, small fluorescent dyes, or photo-controllable groups, the latter enabling photo-regulated translation in vitro and in zebrafish embryos.(34-37) We became particularly interested in the design of cap analogs which could be used as universal reagents for mRNA labeling or modification by the co-transcriptional method. We envisaged that to achieve this, the cap should carry a functional group that enables attachment of various molecules by the same simple chemical method and that the modification site should be selected to minimally interfere with cap-related mRNA functions. As such, in this work we synthesized and characterized a series of cap analogs carrying linkers (spacers) attached to the ribose moiety, which are terminated by amino (NH2) groups enabling site-specific labeling with various labels carrying carboxyl using N-hydroxysuccinimide (NHS) chemistry. We evaluated the analogs as versatile tools for efficient site-specific and sequence-independent labeling of short and full-length mRNAs. We explored two complementary methods to obtain labeled RNA: a co-transcriptional labeling method, in which the label is first attached to a cap analog and then incorporated into the RNA by transcription in vitro, and a post-transcriptional labeling method, in which an NH2-functionalized cap analog is incorporated into the RNA by transcription in vitro followed by chemical labeling of RNA. We also verified if the modified RNA remained translationally active and could be applied in de-capping assays, RNA degradation assays, and RNA visualization in cells. RESULTS m7G cap analogs substituted at the 2′-O or 3′-O positions with long spacers maintain specific interaction with eIF4E and function in translation initiation. Cap analogs functionalized at the 2′-O or 3′-O positions of 7-methylguanosine (m7G), carrying carbamatesubstituted spacers of various lengths terminated by amino (NH2) groups used in this study are shown in Figure 2. We envisaged that due to the reactivity of primary unsubstituted amines towards NHS-esters of carboxylic acids, the novel analogs could give access to capped mRNAs that are fluorescently labeled or otherwise tagged within the 5′ cap and maintain their biological functionality – a concept explored in this work. However, prior to studies on labeling, we performed a few simple biochemical tests on non-labeled cap analogs (Figure 2) to determine the influence of different spacers on biochemical properties of the cap related to translation initiation and, possibly, to select the best candidates for further studies concerning labeling. To this end, we first determined equilibrium association constants of the cap analog complexes with murine translation initiation factor 4E (eIF4E) as well as transcription and translation efficiency in cell lysates (rabbit reticulocyte lysate, RRL).



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 4 of 35

Figure 2. Structures and nomenclature of mRNA 5′ cap analogs and their conjugates studied in this work. A) Reference analogs (1-3): m7GpppG (1) – standard m7G cap analog; m27,3′-O GpppG (2) – anti-reverse cap analog or ARCA, yields mRNAs with translation efficiency superior to 1; ApppG (3) – non-functional cap, yields mRNAs with poor translation efficiency. B) Cap analogs 4-6 carrying NH2 groups to enable mRNA 5′ end labeling: cap with short linker (4), cap with medium linker (5), cap with long linker (6). Label attachment site is marked with Z (for unlabeled analog Z = H). All analogs were synthesized as a mixture of 2′-O and 3′-O regioisomers (ratio of 2′-O:3′-O isomers were as follows: 1:1.79 for 4, 1:1.16 for 5, 1:1.24 for 6); C) Labels (Z) used in experiments: biotin (B), fluorescein (F), and acetyl (Ac). Linkers are colored blue; labels are colored red. Binding to eIF4E. Formation of cap-eIF4E complex is the rate-determining step of cap-dependent translation,(38, 39) therefore high affinity for eIF4E is a desirable feature in the design of cap structures which could confer favorable translational properties to mRNA. The equilibrium association constants (KAS) for complexes of eIF4E with selected analogs were determined using time-synchronized fluorescence quenching titration (Figure 3A, Table S1) and compared to corresponding values for reference cap analogs reported earlier (non-modified cap: m7GpppG (1), commercially available ARCA: m27,3′-OGpppG (2), and non-functional cap: ApppG (3)).(40) Generally, the functionalized analogs were characterized by KAS approximately 1.7–1.9-fold lower than those of m7GpppG, except for analog 6b, for which KAS was 4-fold lower. The decrease in binding could result either from steric effects caused by bulkiness of the carbamate linker or positive charge repulsion between the linker


Page 5 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


and the basic amino acids present in the cap-binding pocket of eIF4E (all NH2 groups are expected to be protonated at pH ~7). Derivatization of analog 4 (4a KAS 7.3 ± 0.1 µM-1 and 4b 6.6 ± 0.1 µM-1 for 2′-O and 3′-O regioisomers, respectively) into uncharged acetyl amide slightly increased the KAS value (4Ac 8.4 ± 0.1 µM-1) bringing it closer to KAS of m27,3′-OGpppG (10.2 µM-1). This suggested that the decrease in affinity for eIF4E observed for amino-functionalized analogs is at least partially caused by repulsion between –NH3+ group from the linker and the protein, and as such, could be alleviated upon linker functionalization.

Figure 3. Biochemical properties of 2′-O and 3′-O substituted cap analogs. A) Association constants (KAS) for complexes of translation initiation factor 4E (eIF4E) and selected 5′ cap analogs (isolated 2′-O and 3′-O isomers, a and b, respectively). Data shown are means from triplicate experiments. B) Relative translation efficiencies of luciferase mRNAs capped with selected analogs (2′-O and 3′-O mixtures or isolated isomers) in rabbit reticulocyte lysate (RRL). Results are color coded according to the type of modification present. Data shown represent means from duplicate experiments. In vitro transcription and translation efficiency. In vitro transcribed (IVT), 5′ capped mRNAs are used as templates for translation of proteins of interest in cellfree systems, cultured cells, and even in whole organisms. Previous studies have shown that modifications of the 5′ cap can influence mRNA translation efficiency significantly.(40) Therefore, we synthesized mRNAs capped with the novel analogs and characterized the transcripts for their translation efficiency to assess the influence of the linker on mRNA functionality. Analogs 1–6 were introduced into mRNAs encoding firefly luciferase in an in vitro transcription reaction under standard conditions.(41) Equivalent amounts of the resulting transcripts were used to program RRL and their overall efficiency of translation after 60 min was determined by luminometry. The results were compared to mRNAs capped with reference analogs 1–3 (m7GpppG, m27,3′-OGpppG, and ApppG). The translation efficiency values for differently capped mRNAs were normalized to the corresponding values for m7GpppG-capped mRNA (RNA-3) (for which translation efficiency was set to 1) and summarized in Figure 3B (for numerical data see Table S1). mRNAs capped with m27,3′-OGpppG and ApppG had translation efficiencies of 1.4 and 0.1, respectively, indicating that the system is well-optimized for cap-dependent translation and reproduces previously reported results.(35) The translation efficiencies for analogs 4–6 varied



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 6 of 35

from 1.1 to 1.7, consistent with all mRNAs being translated in a cap-dependent manner. There was no obvious correlation between translation efficiency and KAS for eIF4E. Instead, an increasing trend in translation efficiency was observed with shortening the linker. Interestingly, translation efficiency for mRNA capped with a charge-masked analog carrying the shortest linker 4Ac (1.14 ± 0.3) was notably lower than the translation efficiency for unmasked analog 4 (1.66 ± 0.15). Overall, the translational efficiencies for analogs 4–6 were sufficient to consider them all as reagents for mRNA modification. Novel cap analogs provide access to fluorescently labeled and biotinylated RNAs that are translationally active by co- and post-transcriptional labeling approaches. Co-transcriptional versus post-transcriptional labeling approach. Most of the covalent RNA labeling methods can be classified according to the relative order of RNA synthesis and labeling reactions. Incorporation of a fluorescent nucleotide derivative into mRNA during in vitro transcription is termed co-transcriptional labeling, while incorporation of a functionalized nucleotide during transcription and its subsequent labeling by chemical reaction is termed post-transcriptional labeling. We envisaged that, in the context of mRNA cap labeling using NHS chemistry, both of these approaches, schematically depicted in Figure 4A, are of potential utility. Consequently, in this work we concurrently explored both of these to define their scope and limitations. The conditions for reactions with active NHS esters were optimized for NH2-functionalized dinucleotides and RNAs of various lengths and sequences. Next, we compared the co- and post-transcriptional strategies by testing the quality and functionality of the resulting transcripts, level of non-specific labeling, and the functionality of labels in the resulting conjugates.

Figure 4. A) Two approaches to mRNA 5′ cap labeling explored in this work: co-transcriptional (co-trx) and post-transcriptional (post-trx) labeling; B) NHS-labeling of amino-functionalized cap analog; Label: 5(6)carboxyfluorescein,

biotin

or

acetic

acid;

TSTU:

N,N,N’,N’-Tetramethyl-O-(N-succinimidyl)uronium

tetrafluoroborate Fluorescent labeling of dinucleotide cap structures. The labeling reactions were first optimized for dinucleotide analogs 4–6. Labeling was performed by mixing aqueous solutions of cap analogs, borate buffered to pH 8.5, with DMSO solution of a compound carrying NHSactivated carboxyl group (carboxyfluorescein, biotin, or acetic acid; Figure 4B). The conversion of carboxylic acids into NHS-esters was performed in situ by incubation with TSTU and triethylamine for 30 min at room


Page 7 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


temperature.(42) The progress of labeling reactions was monitored by HPLC. Typically, a 2–4 molar excess of NHS ester was required to achieve complete labeling of the cap analog within 1 h. We observed high selectivity of NHS-esters towards primary amino groups terminating the linker moieties despite the presence of many other functional groups in the cap structure. Comparing reactivity between particular analogs, we found that the analog with the shortest linker (4), reacted less efficiently than analogs 5 and 6. Importantly, reagents such as ammonium cations or amino-containing buffering reagents (e.g., Tris) undergo side reactions with the NHSesters and, as such, their presence in labeling reaction mixtures should be avoided. The labeled products 4Ac, 5F, 5B, 6aF, 6B were isolated by semi-preparative HPLC directly from diluted reaction mixtures, before their use to test co-transcriptional labeling of RNA. Synthesis and properties of co-transcriptionally labeled RNAs. Labeled cap analogs (5F, 5B, 6aF, 6B) were tested as reagents for the synthesis of capped mRNAs. The transcription reactions yielding differently capped firefly luciferase mRNAs (RNA-3) and the translation experiments in RRL for those mRNAs were carried out under the same conditions as those described above for the corresponding unlabeled caps. For all co-transcriptionally labeled RNAs we found translation efficiency to be in the range of 0.48 to 1.15, which is from 15 to 64% lower than that for RNAs with corresponding unlabeled cap analogs (Table 1). The presence of fluorescence label in the transcript capped with analog 5F was confirmed by fluorescence emission spectroscopy. Concentration-dependent measurements revealed that the fluorescence emission could be detected at mRNA concentrations above 1 nM (0.6 ng/µl) (Figure S1A), which is about 10fold lower than the detection limit of a labeled dinucleotide (Figure S1B) and consistent with the presence of no more than one fluorescent label per mRNA molecule. Table 1. Relative translation efficiencies in rabbit reticulocyte lysate for luciferase mRNAs (RNAs-3) cotranscriptionally capped with labeled cap analogs 5F, 5B and 6F or reference analogs 1, 2, 5, 6. Cap analog at the mRNA 5′ enda

Relative translation efficiencyb

Cap analog at the mRNA 5′ end

Relative translation efficiencyb

1 (m7GpppG)

1

2 (m27,3′-OGpppG)

1.46 ± 0.14

5F

0.48 ± 0.03

5B

1.15 ± 0.23

5

1.35 ± 0.18

6B

0.98 ± 0.04

6

1.09 ± 0.01

a

b

Abbreviations: F: fluorescein (FAM), B: biotin; Data shown are means ± S.D. from duplicate experiments.

The reason for lowered translation efficiency of labeled mRNAs could be twofold. First, the efficiency of incorporation of cap moieties during in vitro transcription (capping efficiency) could be lower for labeled caps compared to unlabeled ones, resulting in a smaller fraction of translationally active mRNAs in the transcription product. Second, the label could diminish affinity of the cap for the eIF4E because of steric reasons, in turn decreasing the translational activity of the mRNA. However, the second option appeared to be less likely due to the large distance between the label and cap moiety ensured by the presence of the 6- or 13-atom linkers. To estimate capping efficiency achieved with cap analogs 5 and 6 we performed transcription reactions yielding short RNAs (RNA-1), which were analyzed by electrophoresis under conditions enabling resolution of capped and uncapped transcripts (Figure S2). We found that while unlabeled dinucleotides 5 and 6 are incorporated into



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 8 of 35

RNA with efficiency comparable to m7GpppG (53–77% of capping), their labeled versions are less efficient capping reagents (2–23% of capping). This suggested that the lower translational efficiencies of mRNAs capped with compounds 5F, 5B, and 6B arise, to a large extent, from lower capping efficiency compared to 5 and 6. Nonetheless, the capping efficiencies determined for short transcripts (RNA-1) may not be perfect estimates for capping efficiencies of longer transcripts, since in the latter case the cap analog to NTP ratio increases considerably more rapidly with the progress of the in vitro transcription reaction. These observations turned our attention to the concept of post-transcriptional RNA labeling. As such, different RNAs co-transcriptionally capped with 5 and 6 were tested as substrates in post-transcriptional labeling reactions with fluorescein and biotin NHS esters. Post-transcriptional fluorescent labeling of short and long transcripts. Fluorescent labeling was first optimized on short, 24 nt RNA-1 transcripts capped with 6. The conditions for labeling reactions were, essentially similar to those used for labeled dinucleotides. However, optimization of active ester excess over RNA was required to achieve sufficient labeling within a reasonable time (~1 h) and to minimize non-specific (random) labeling. The transcripts were incubated in aqueous borate buffer pH 8.5 with DMSO solution of NHS ester of fluorescein (FAM-NHS; prepared in situ), followed by analysis by PAGE under conditions enabling resolution of uncapped, capped and labeled RNA (Figure 5A). To determine the extent of non-specific labeling, uncapped transcripts and transcripts capped with analog 1 (m7GpppG) were subjected to the same procedure. The electrophoretic analysis was performed first with fluorescein visualization (emission ≥ 510 nm) followed by staining and visualization of total RNA. Upon initial trials, we found that the optimal labeling of RNA-1 capped with 6 is achieved by treatment with 100-fold excess of FAM-NHS for 1 h (Figure S3A, B). The densitometric analysis revealed that around 60% of RNA-1 capped with 6 was labeled under those conditions. Moreover, the labeling reaction was highly specific since uncapped RNA and RNA capped with m7GpppG did not exhibit FAM emission after FAM-NHS treatment (Figure 5A). Next, we moved to labeling full-length transcripts encoding firefly luciferase (RNA-4; 1.6 kb) capped with analog 6 and labeled with in situ generated FAM-NHS. The labeling was very inefficient under conditions employed for 24 nt RNA-1. Upon optimization, we found that efficient labeling of RNA-4 is achieved in the presence of 10,000-fold excess of FAM-NHS (Figure S3C). However, under those conditions uncapped or m7GpppG-capped transcripts were found to also undergo (non-specific) labeling (Figure 5B). The level of the non-specific labeling was estimated to be as high as 25%. We envisaged that the non-specific labeling may result from the use of in situ generated FAM-NHS, as it contained residual activating reagents, which could be the cause of side-reactions when used in such high excess. Therefore, we next compared the efficiency and specificity of labeling using either in situ activated FAM-NHS or purified FAM-NHS (from a commercial source). Densitometric analysis revealed that the use of purified FAM-NHS decreased the non-specific labeling to 8% (Figure 5B). Finally, we prepared RNA-3 transcripts labeled by the post-transcriptional approach and tested their efficiency of translation in RRLs (Table 2). The biotinylated transcripts were translated at efficiencies similar to transcripts labeled via the co-transcriptional approach, whereas the fluorescein-labeled transcripts had higher translation efficiencies than co-transcriptionally labeled ones.


Page 9 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Table 2. Relative translation efficiencies in rabbit reticulocyte lysate for luciferase mRNAs (RNAs-3) capped with cap analog 5 and labeled post-transcriptionally with fluorescein or biotin along with the same data for unlabeled reference mRNAs (uncapped and capped with 1, 2, 3, or 5). Reference mRNAs Unlabeled mRNA

Relative translation efficiencya

Unlabeled mRNA


1-RNA-3

1

2-RNA-3

1.56 ± 0.02

3-RNA-3

0.11 ± 0.04

pppG-RNA-3 (uncapped)

0.10 ± 0.02

5-RNA-3

1.46 ± 0.10 Post-trx labeled mRNAs

Substrate mRNA

Labeling reagent

Labeled mRNAb


5-RNA-3

FAM-NHS

5F-RNA-3 (post-trx)

0.96 ± 0.29

5-RNA-3

Biotin-NHS

5B-RNA-3 (post-trx)

1.13 ± 0.26

a

b

Data shown are means ± S.D. from 3 to 5 experiments. Abbreviations: F: fluorescein (FAM), B: biotin;

RNA biotinylation. Another label tested in this study was D-biotin, which has found many interesting applications in biotechnology, owing to its extremely high affinity to (strept)avidin. In order to verify the labeling efficiency as well as the functionality of label conjugated with RNA, we prepared 140-nt transcripts (RNA-2) biotinylated by both coand post-transcriptional protocol and used an electrophoretic mobility shift assay (EMSA) in a non-denaturing agarose gel to monitor binding to recombinant tetrameric streptavidin (Figure 5C). In the presence of streptavidin, a fraction of RNA migrated distinctly slower than in a pool containing no streptavidin, clearly illustrating formation of the complex. However, even in mixtures containing a large excess of streptavidin, the faster migrating band of non-bound RNA was still observed, consistent with the presence of a fraction of transcripts that are unlabeled or uncapped. The fractions of biotinylated RNA products obtained by co- and posttranscriptional procedures were similar and constituted 60–70% of total RNA (by densitometry), indicating higher capping efficiency of RNA-2 compared to that observed for short (24 nt) RNA-1.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 10 of 35

Figure 5. Demonstration of post-transcriptional labeling procedures for RNAs of various lengths. A) Electrophoretic analysis of post-transcriptionally FAM-labeled 24-nt RNA-1 capped with cap 6a (labeling conditions: 1 h, 100-fold excess of FAM-NHS obtained by in situ activation). Uncapped and m7GpppG-capped RNAs were used as negative controls. B) Electrophoretic analysis of post-transcriptionally FAM-labeled luciferase mRNAs (RNA-4) capped with cap 6 (labeling conditions: 1 h, 10,000-fold excess of FAM-NHS obtained either by in situ activation or from a commercial source). Uncapped RNA and RNAs cotranscriptionally capped with m7GpppG and cap 5F were used as controls. Whole gel images are shown in the Supporting Information (Figure S4). C) Electrophoretic mobility shift assay showing formation of the complexes of streptavidin and 140-nt RNAs (RNA-2) biotinylated within 5′ cap structures by either the co-transcriptional approach (transcription reaction in the presence of cap 5B) or the post-transcriptional approach (reaction of 5BRNA-2 with NHS-biotin). 1-RNA-2 was used as a negative control. 5 ′ -Fluorescently labeled and capped RNAs are suitable for RNA degradation assays involving recombinant proteins and in cell extracts. Assaying decapping activity. mRNA 5′-to-3′ degradation is one of the key pathways regulating gene expression. The pathway is initiated by Dcp1-Dcp2, which has been identified as the major mRNA decapping enzyme. Dcp1-Dcp2 cleaves the cap between α and β phosphates to release m7GDP and 5′-monophosphorylated RNA exposed to 5′-exonucleases. Other enzymes from the Nudix (nucleoside diphosphate linked to another moiety X) family have recently been shown to have decapping activity.(43, 44) Decapping activity is often assayed in vitro using oligonucleotides radioactively [32P]-labeled within the γ-phosphate of the cap and analysis of the reaction products by electrophoresis followed autoradiography. Upon decapping, α-[32P]-m7GDP molecules are released, leading to decreases in radioactivity of the total RNA pool. We envisaged that our fluorescently labeled, capped RNAs could be a viable alternative to [32P]-labeling in the context of such applications. To verify this, we used short transcripts (RNA-1) fluorescently labeled using either post-transcriptional or co-transcriptional labeling protocols, incubated them with Dcp1-Dcp2 complex, analyzed the products by gel electrophoresis, and visualized for fluorescein emission, followed by total RNA staining (Figure 6A). Visualization of total RNA staining revealed that all capped transcripts (labeled and unlabeled) were completely hydrolyzed by the enzyme within 1 h. Visualization of fluorescein emission revealed complete disappearance of FAM-RNA related bands in the lanes corresponding to RNAs treated with Dcp1-Dcp2 proving that incorporation of the label into ribose of 7-methylguanosine does not prevent decapping. The result also confirms that the observed fluorescence is a result of highly specific labeling of the RNA within the caps’ 7-methylguanosine, which is cleaved off by the enzyme as a labeled diphosphate (FAM-m7GDP) leaving unlabeled RNA. Next, we performed an analogous decapping experiment using Dcp1-Dcp2 on full-length mRNAs (RNA-4) and analyzed the samples by electrophoresis. Fluorescein visualization confirmed that decapping occurs both on post-transcriptionally and co-transcriptionally labeled mRNA (Figure 6B). For co-transcriptionally labeled mRNA, the fluorescent signal of 5F-RNA-4 vanished completely upon decapping, confirming that the fluorescent label was placed exclusively within the 5′ cap. For post-transcriptionally labeled 6a-RNA-4, residual RNA fluorescence of intensity around 30 % (in situ FAM-NHS) and 15 % (commercial FAM-NHS) of the initial fluorescence was still observed after decapping. These values are consistent with the level of non-specific labeling determined upon optimization of labeling reactions (Figure 5B). Control mRNAs either uncapped or capped with m7GpppG after being subjected to post-transcriptional labeling procedure had fluorescence


Page 11 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


intensities similar to decapped transcript 6a-RNA-4. The fluorescence of control transcripts did not disappear upon incubation with Dcp1-Dcp2, additionally confirming that this residual fluorescence is related to nonspecific labeling of mRNAs. To confirm that the observed changes in fluorescence intensity are associated with decapping, we treated the post-decapping mixtures with Xrn1, which is a 5′-exoribonuclease that readily hydrolyses 5′-monophosphorylated RNA, but is inactive towards 5′-capped RNAs. Total RNA staining confirmed that only RNAs post Dcp1-Dcp2 treatment are readily hydrolyzed by Xrn1. Finally, using a posttranscriptionally labeled 5-RNA-3 we performed a decapping experiment for hNudt16, which is another Nudix family enzyme with in vitro activity similar to Dcp1-Dcp2. Electrophoretic analysis of the reaction mixtures at different time points (0, 2, 30 and 60 min) revealed changes in fluorescein emission from RNA consistent with progressive decapping over time (Figure S5).

Figure 6. Utility of fluorescently labeled transcripts in RNA decapping assays. A) Dcp2/Dcp1-catalyzed decapping of RNA-1 (24 nt) obtained by either a post- or co-transcriptional protocol. B) Electrophoretic analysis of post-transcriptionally labeled luciferase mRNAs before and after incubation with decapping enzyme SpDcp1Dcp2 followed by incubation with Xrn1. Gels were visualized both in fluorescence mode and by SYBR® Gold or EtBr staining; mRNAs co-transcriptionally labeled with 5F were used as a control. Whole gel images are shown in Supporting information (Figure S6). Monitoring RNA 3′-to-5′ degradation in HeLa extract. We next studied if cap-labeled RNAs can be used as probes to monitor RNA degradation in a more complex medium, such as cellular extract. To this end, transcript 6-RNA-1a co- or post-transcriptionally labeled with fluorescein was incubated in a cytoplasmic extract from HeLa cells. Electrophoretic analysis of samples collected from the reaction mixture at different time points revealed decay of the starting fluorescent material and increasing fluorescence intensity in faster migrating bands (Figure 7). With increasing time, the majority of fluorescence accumulated in a product that migrated at the same height as FAM-labeled cap 6, which is the expected product of complete 3′-to-5′ RNA degradation. To confirm that the observed degradation is catalyzed by 3′-nucleases present in HeLa cells, we incubated the same RNA with heat-denatured extract and observed no such reaction.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 12 of 35

Figure 7. Utility of fluorescently labeled transcripts in monitoring of 3′-to-5′ RNA decay. Electrophoretic analysis of RNA-1a (35 nt) labeled by a post- (A) and co-transcriptional (B) protocol incubated with HeLa cytoplasmic extract for the indicated times. Heat-denatured extract (–) was used as a negative control. Lanes with fluorescently labeled cap (6aF, 5F) are shown for reference. 5′-Fluorescently labeled and capped mRNAs are amenable to detection and undergo translation in living cells. Finally, we studied the properties of our labeled mRNAs in living cells. We first attempted detection of the labeled mRNAs inside cells. Cultured HeLa cells were transfected for 5 h with firefly luciferase mRNA (RNA-5) terminated by a 128 nt 3′-polyA tail and carrying cap 6 or cap 5 labeled by the post- or co-transcriptional approach, respectively. Moreover, RNA-5 capped with unlabeled analog 6 served as a negative control. The cells were then fixed and visualized by confocal microscopy (Figure 8). The FAM fluorescence signal was detected in cells that were transfected with mRNA carrying fluorescently labeled cap 6 and 5, but not in untreated cells or cells treated with mRNA capped with unlabeled cap analog 6. Similar fluorescence intensities were observed for co- and post-transcriptionally labeled mRNAs, and in both cases, a notably lower fluorescence intensity was observed inside the nucleus, in agreement with the cytoplasmic localization of mRNA.


Page 13 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 8. Localization of fluorescently labeled mRNA in HeLa cells. HeLa cells were transfected with cotranscriptionally (5F-RNA-5 co-trx) or post-transcriptionally (6aF-RNA-5 post-trx) labeled mRNA encoding firefly luciferase. HeLa cells transfected with mRNA capped with unlabeled analog 6a (6a-RNA-5) and mocktreated cells were used as negative controls. Panels from left to right show fluorescein fluorescence, Hoechst fluorescence, merge of these two panels (merge 1), bright field and merge of all images. Scale bars are 20 µm. Finally, we also verified if the transfected labeled mRNAs are translationally active in the cells. The translation efficiencies were determined for the same firefly luciferase mRNA sequence (RNA-5) carrying cap 6, cap 6 subjected to post-transcriptional labeling, cap 5F (yielding co-transcriptionally labeled RNA), cap analogs 2 and 3 (translationally active and non-active reference mRNAs, respectively), and uncapped mRNA (pppG-RNA-5). To eliminate errors arising from random differences in transfection efficiency, mRNA encoding Renilla luciferase and capped with cap analog 2 (m27,3′-OGpppG) was included in each experiment as an internal reference and control. HeLa cells were transfected for 5 h with a 7:3 mixture of RNA-5 and control mRNA, followed by cell lysis and determination of firefly luciferase and Renilla luciferase activities by luminometry. The raw firefly luciferase activity values were normalized to the values obtained for the internal control (Renilla luciferase activity), as described in the experimental section. The normalized firefly luciferase activity values for differently capped RNAs-5 are shown in Figure 9. The results indicated that the labeled transcripts obtained by both co- and post-transcriptional approaches undergo translation in living cells, albeit less efficiently than their non-labeled counterparts. Post-transcriptionally labeled mRNA (6aF-RNA-5) had translational activity approximately 40% lower than its unlabeled version (6a-RNA-5). Nonetheless, both transcripts were translated in a cap-dependent manner, since they underwent translation at least 60 times more efficiently than mRNA carrying the non-functional cap structure 3 or uncapped mRNA (pppG-RNA-5). The difference between labeled



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 14 of 35

and unlabeled mRNA (6aF-RNA-5 versus 6a-RNA-5) most likely resulted directly from the influence of the presence of the fluorescent label, since the labeling procedure itself (FAM-NHS treatment) had no effect on translation efficiency, as seen from comparison of translation efficiency for mRNA capped with cap analog 2 with the same mRNA post NHS-treatment (Fig.9, 2 versus 2 + F). Fluorescently labeled mRNA (5F-RNA-5) obtained by the co-transcriptional procedure was translated slightly less efficiently than post-transcriptionally prepared 6aF-RNA-5.

Figure 9. Translation efficiencies in HeLa cells for differently capped firefly luciferase mRNAs (RNAs-5). Cells were transfected for 5 h with a mixture of 70 ng luciferase mRNA (RNA-5), uncapped or carrying the studied cap structure, and 30 ng control Renilla luciferase mRNA capped with analog 2 (2-RNARluc). The activity of both luciferases was measured using a dual-reporter assay, followed by normalization of firefly luciferase activity to Renilla activity to account for transfection efficiency differences as described in the experimental section. “2”, “3”, “6a”, “5F” denote RNA-5 transcripts co-transcriptionally capped at their 5′ ends with cap analogs 2, 3, 6a, 5F, respectively. “pppG” denotes uncapped (GTP-initiated) RNA-5, while “2+F” and “6a+F” refer to RNA-5 capped with analogs 2 and 6a, respectively, subjected to post-transcriptional labeling procedure with commercial FAM-NHS. “Mock” refers to cells transfected only with control RNA (2-RNARluc). Data shown are average values from triplicate experiments ± S.D. Values above each bar represent mean normalized luciferase activity. DISCUSSION In this work, we aimed to develop new mRNA 5′ cap analogs that could be useful as reagents for site-specific RNA labeling, thereby providing novel tools to investigate mRNA, especially in the context of cap-dependent processes. We envisaged that to develop a method accessible to the broad biological community, the cap labeling approach should be based on a well-established chemical method that is reliable, reproducible, adaptable to a variety of fluorescent dyes and biological tags, and easy to perform without specialized equipment. Therefore,


Page 15 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


we turned our attention to N-hydroxysuccinimide (NHS) chemistry, which is one of the most popular methods used for labeling and conjugation of biomolecules, and synthesized a set of NH2-functionalized cap analogs that are susceptible to labeling with NHS-esters of fluorescein and biotin (Figures 2 and 3). This approach offers a great deal of flexibility regarding tag selection, since almost every fluorescent dye or biological tag is commercially available in the form of a carboxylic acid derivative. The carboxylic acids can be easily converted into NHS esters using in situ activation procedures or potentially any other activation procedure adopted from peptide chemistry (such as carbodiimide or triazolol derivatization); many of these are also commercially available in the NHS-activated form. The high selectivity of the NHS-labeling approach allowed us to independently test two labeling strategies, which we termed co-transcriptional and post-transcriptional labeling (Figure 4). The co-transcriptional approach relied on one-step incorporation of a chemically synthesized labeled cap analog into RNA by in vitro transcription, i.e., a typical procedure applied for preparation of capped RNAs.(45, 46) The post-transcriptional approach encompassed two steps: first, co-transcriptional incorporation of a NH2-functionalized cap into RNA and second, chemical labeling of the resulting transcript. We found both approaches to be suitable for labeling of short and long RNAs, although each had different advantages and limitations. The main advantage of co-transcriptional labeling was simplicity and excellent specificity (i.e., only a very small degree of non-specific labeling). Unfortunately, the efficiency of incorporation of labeled dinucleotides into short RNAs (% of capping) was rather low (~20 % for biotin and ~2 % for fluorescein) as compared to capping efficiencies for unlabeled caps (50–80%) (Figure S2). Nonetheless, capping with labeled analogs appeared to be more efficient for longer RNAs, as can be noted from direct comparisons (e.g., biotinylated RNA fractions in Figure S2 and Figure 5C) and indirectly from similar biochemical properties of co- and post-transcriptionally labeled full-length mRNAs (e.g., see Tables 2 and 3, Figure 9). Although the redundant, uncapped RNA should not interfere with many typical biochemical applications such as decapping assays and RNA decay assays, if necessary, it can be removed from the RNA batch by treatment with 5′ polyphosphatase and 5′-exonuclease (e.g., Xrn1). The main advantage of the post-transcriptional labeling method was that it enabled to overcome the problem of inefficient capping, observed especially for short RNAs. The conditions for reaction with active NHS esters were successfully optimized for RNAs of various lengths and sequences. Overall, we found that up to 100-fold higher excess of labeling reagent was necessary for efficient labeling of long RNAs compared to short ones. The labeling of short transcripts was highly specific for the 5′ cap, whereas for longer transcripts non-specific labeling of the mRNA was observed at 8–30% (better results were obtained for purified NHS-ester than for an in situ generated one). Among numerous approaches developed for mRNA cap labeling, most desirable are the ones that do not alter the biological activity of mRNA. Therefore, we studied how the proposed RNA labeling strategy influences translation and decapping. Labeled caps were less efficient initiators of cap-dependent translation than were unlabeled parent compounds, both in cell lysates (RRLs) and in cultured HeLa cells (Tables 2 and 3, Figure 9). Nevertheless, the labeled mRNAs were translated in a cap-dependent manner as they were translated notably more efficiently than uncapped mRNA (pppG-RNA) or mRNA carrying the non-functional cap analog 3



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 16 of 35

(ApppG-RNA). Decreases in translational activity of labeled mRNAs could be attributed to two reasons: (i) inefficient incorporation of the labeled cap analog into mRNA (only relevant for co-transcriptional labeling) (ii) the direct influence of label moiety on mRNA translation efficiency (relevant for both co-transcriptional and post-transcriptional labeling). Both factors appear to contribute to the decrease in translation efficiency of labeled mRNAs, especially in the case of the more bulky fluorescein label (as opposed to smaller biotin moieties). We also showed that fluorescein-labeled transcripts enable observation of decapping by Dcp1-Dcp2 and hNudt16 in vitro and RNA decay in HeLa cell extracts, thereby demonstrating utility in studies of both 5′-to3′ and to 3′-to-5′ RNA decay. However, especially in the context of 5′-initiated degradation, one should keep in mind that the presence of the label may influence the kinetics of decapping processes catalyzed by different proteins to various extents. Finally, we demonstrated that the transcripts, labeled either by co-transcriptional or post-transcriptional approaches, can be detected in cultured cells post transfection. In conclusion, we demonstrated that both co- and post-transcriptionally labeled RNAs are compatible with assaying decapping enzyme activity, can be committed to RNA degradation pathways, can be visualized in cells and undergo cap-dependent translation in vitro and in vivo. Therefore, we envisage that the reported labeling approaches not only provide alternatives to radioactive mRNA cap labeling, but also open avenues for novel, and so far unexplored, applications. Further studies on optimization of the capping efficiencies and enhancing translational efficiencies of modified mRNAs are in progress. EXPERIMENTAL PROCEDURES MATERIALS Solvents, chemical reagents, and labels (D-biotin, carboxyfluorescein) were purchased from Sigma Aldrich and used without further treatment. For fluorescent labeling, a mixture of 5/6-carboxyfluorescein was used. The regioisomers of 5(6)-carboxyfluorescein conjugates were separated by RP-HPLC (assigned with F1 – isomer 6 or F2 – isomer 5 suffix, according to the elution order) and characterized separately. General information on the chemical synthesis, including reaction conditions, purification protocols and products characterization are available in Supplementary Information. METHODS Preparation of recombinant proteins (eIF4E and SpDcp1-Dcp2). Mouse eIF4E (residues 28–217) was expressed in Escherichia coli strain BL21(DE3) as inclusion bodies. Guanidinium-solubilized protein was refolded by one-step dialysis, and purified by ion-exchange chromatography on HiTrap SP column (GE Healthcare) without exposure to cap analogs. The concentration of eIF4E was determined spectrophotometrically (ε280nm = 52,940 L·mol-1cm-1).(47) Schizosaccharomyces pombe Dcp1-Dcp2 complex was prepared as described previously.(48) Cap-eIF4E association constants. Association constants for eIF4E-cap analog complexes were determined by intrinsic tryptophan fluorescence quenching titration. Measurements were carried out on an LS-55 spectrofluorometer (Perkin Elmer Co.) in 50 mM HEPES/KOH (pH 7.2), 100 mM KCl, 0.5 mM EDTA, 1 mM DTT at 20.0 ± 0.3°C. Aliquots of 1 µl


Page 17 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


increasing concentration of cap analog solutions were added to 1.4 ml of 0.1 µM mouse eIF4E(28-217) protein solutions. Fluorescence intensities (excitation at 280 nm with 2.5 nm bandwidth and detection at 340 nm with 4 nm bandwidth and 290 nm cut-off filter) were corrected taking into account sample dilution and the inner filter effect. Equilibrium association constants (KAS) were determined by fitting the theoretical dependence of the fluorescence intensity on the total concentration of cap analog to the experimental data points according to an equation described previously.(49) The final KAS were calculated as weighted averages of three to five independent titration experiments. Numerical nonlinear least-squares regression analysis was performed using OriginPro8 (Microcal Software Inc., USA). Labeling of cap analogs. Amino- or carboxy-functionalized dinucleotide cap analogs were labeled using NHS-activation chemistry, as described before.(42) Active NHS esters were generated in situ by addition of equimolar amounts of TSTU to a 0.4 M solution of appropriate carboxylic acids (either cap analog or label) in DMSO alkalized with 2-fold molar excess of Et3N. Prepared NHS-ester (2–4 molar excess) was added portionwise to a solution of an aminofunctionalized cap analog in borate buffer pH 8.5. Progress of the reaction was monitored using RP HPLC and the products were isolated from diluted reaction mixtures by semi-preparative RP HPLC. The detailed protocols for labeling of cap analogs, including reagents amount and preparative yields are described in Supplementary Information. RNA synthesis. In order to perform all presented experiments, RNAs with different sequence composition and length were synthetized by means of in vitro transcription reactions (Table 3). A detailed description of synthesis of all RNAs used in this study is presented below. Table 3. Definition of RNA abbreviations used in this study. No.

RNA lengtha

RNA sequence or sequence description

Type of experiment

Poly(A)

RNA-1a

35

GA AGAAGC GGGCAU GCGGCC AGCCAU AGCCGA UCA

RNA degradation in HeLa extract (Fig. 7)

˗

RNA-1

25

GA AGAAGC GGGCAU GCGGCC AGCCA

short RNA labeling (Fig. 5A, S3A-B); RNA decapping assay (Fig. 6A); RNA capping efficiency (Fig. S2)

˗

RNA-2

140

β-globin 5′UTR + 97 nts of luciferase gene

EMSA (Fig. 5C); Nudt16 assay (Fig. S5)

˗

RNA-3

1820

β-globin 5′UTR + luciferase gene + A31 + N49

translation in RRL (Fig. 3, Table 1, 2)

+

RNA-4

1676 Firefly luciferase gene

mRNA labeling (Fig. 5B, S3C), RNA decapping assay (Fig. 6B)

˗

RNA-5

2105

Firefly luciferase gene + 2 repeats of β-globin 3′ UTR + A128

mRNA localization (Fig. 8); translation HeLa cells (Fig. 9)

+

2-RNARluc

1448

Renilla luciferase gene + 2 repeats of β-globin 3′ UTR + A128

translation in HeLa cells (Fig. 9)

+

a

The lengths refer to uncapped RNAs, corresponding capped RNAs are one nucleotide longer due to m7G

RNA-1a



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 18 of 35

RNAs-1a carrying various cap structures (labeled or unlabeled, as specified in sections below) were 35 nt precursors to the synthesis of RNAs-1. RNA-1a was also used in degradation experiments in HeLa cell extracts. In vitro transcription reaction mixture (20 µl) was incubated at 40°C for 2 h and contained RNA Pol Buffer (New England BioLabs), 1 U/µl SP6 polymerase (ThermoFisher Scientific), 1 U/µl RiboLock RNase Inhibitor (ThermoFisher Scientific), 0.5 mM ATP/CTP/UTP, 0.125 mM GTP, 1.25 mM cap analog, and 0.1 µM annealed oligonucleotides (ATACGATTTAGGTGACACTATAGAAGAAGCGGGCATGCGGCCAGCCATAGCCGATCA TGATCGGCTATGGCTGGCCGCATGCCCGCTTCTTCTATAGTGTCACCTAAATCGTAT)

and as

template.

After 2 h of incubation, 1 U/µl DNase I (Ambion) was added and incubation was continued for 30 min at 37°C followed by addition of EDTA to a final concentration of 25 µM. RNA-1 Differently capped RNAs-1 were substrates used in a Dcp2 decapping assay. To improve the homogeneity of synthesized short RNAs-1a, the transcripts (at 1 µM) were incubated with 1 µM DNAzyme 10–23 (TGATCGGCTAGGCTAGCTACAACGAGGCTGGCCGC) in 50 mM MgCl2 and 50 mM Tris-HCl pH 8.0 for 1 h at 37°C (Coleman et al., 2004), which produced 3′ end-trimmed RNAs (RNA-1). Following incubation, nucleic acids were ethanol-precipitated; the obtained pellet was dissolved in deionized water and treated with 1 U/µl DNase I for 30 min at 37°C. After that, RNAs were phenol/chloroform extracted, ethanol precipitated and the recovered pellet was dissolved in deionized water. RNA-2 RNAs-2 were substrates used in RNA biotinylation reactions and electrophoretic mobility shifts assay with streptavidin. 140-nt long transcripts carrying cap 5 or its biotinylated derivative 5B were synthetized by in vitro transcription with SP6 polymerase as described previously.(35) Typical in vitro transcription reaction (40 µl) contained: RNA Pol Buffer, 0.7 µg of DNA template, 1 U/µl RiboLock RNase Inhibitor, 0.5 mM ATP/CTP/UTP, 0.1 mM GTP, and 0.5 mM cap analog. The reaction mixture was preincubated at 37°C for 5 minutes before addition of SP6 RNA polymerase (final concentration 1 U/µl) and the reaction was continued for 45 minutes at 37°C. Following transcription, 1 U/µL DNase RQ1 (Promega) was added and incubation was continued for 20 min at 37°C. RNA-3 RNAs-3 carrying various cap structures were used in translation experiments performed in RRL and hNudt16catalyzed decapping reactions. Capped mRNAs encoding firefly luciferase were synthetized by in vitro transcription with SP6 polymerase as described previously.(50) In brief, the PCR product was generated based on the pSP-rbBluc plasmid,(51) with an introduced sequence of 31 As at the EcoRI site using primers (ATTTAGGTGACACTATAG and GTAATACGACTCACTATAGGG) that allowed the introduction of promoter sequences for SP6 polymerase. Typical in vitro transcription reactions (40 µl) contained: SP6 transcription buffer, 0.7 µg of DNA template, 1 U/µl RiboLock RNase Inhibitor, 0.5 mM ATP/CTP/UTP and 0.1 mM GTP and 0.5 mM cap analog. The reaction mixture was preincubated at 37°C for 5 minutes before addition


Page 19 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


of SP6 RNA polymerase and the reaction was continued for 45 minutes at 37°C. Following transcription, 1 U/µL DNase RQ1 (Promega) was added and incubation was continued for 20 min at 37°C. RNA-4 RNA-4 was used in Dcp2/Xrn1 degradation assays. mRNA encoding firefly luciferase was synthetized using PCR

product

generated

from

plasmid

pGEM-luc

(Promega)

using

(ATTTAGGTGACACTATAGAAGTACTGTTGGTAAAGCCACCATGGAAGACGCCAAAAACAT

primers and

TTACAATTTGGACTTTCCGCCCT) that allow the introduction of promoter sequences for SP6 polymerase as a template. Typical in vitro transcription reactions (20 µl) were incubated at 40°C for 2 h and contained: RNA Pol Buffer, 1 U/µl SP6 polymerase, 1 U/µl RiboLock RNase Inhibitor, 0.5 mM ATP/CTP/UTP, 0.125 mM GTP, 1.25 mM cap analog and 5 µg/µl PCR product as a template. Following 2 h incubation, 1 U/µl DNase I was added and incubation was continued for 30 min at 37°C, after which EDTA was added to 25 µM final concentration. RNA-5 RNA-5 was used in translation and visualization experiments performed in HeLa cells. Capped mRNAs encoding firefly luciferase and a 128 nt poly(A) tail were synthetized by in vitro transcription with SP6 polymerase. pJET_luc_128A plasmid digested with AarI (ThermoFisher Scientific) was used as a template. This plasmid was obtained by cloning the cDNA sequence encoding firefly luciferase, two repeats of β-globin 3′UTR and 128 adenines from hRLuc-pRNA2(A)128 plasmid (52) into pJET1.2 vector (ThermoFisher Scientific). Typical in vitro transcription reactions (20 µl) were incubated at 40°C for 2 h and contained: 1 U/µl SP6 polymerase, 1 U/µl RiboLock RNase Inhibitor, 0.5 mM ATP/CTP/UTP, 0.125 mM GTP, 1.25 mM cap analog and 5 µg/µl digested plasmid as template. Following 2 h incubation, 1 U/µl DNase I was added and incubation was continued for 30 min at 37°C. EDTA then was added to 25 µM final concentration. Control mRNA for translation efficiency experiment in HeLa cells (2-RNARluc) 2-RNARluc transcript was used as a transfection efficiency control in translation efficiency experiments in HeLa cells. mRNA encoding Renilla luciferase, carrying a 128 nt poly(A) tail, and capped with compound 2 (m27,3′O

GpppG) at the 5′ end was synthetized by in vitro transcription with T7 polymerase (New England BioLabs).

Typical in vitro transcription reactions (20 µl) were incubated at 37°C for 2 h and contained: 1 U/µl T7 polymerase, 1 U/µl RiboLock RNase Inhibitor, 0.5 mM ATP/CTP/UTP, 0.125 mM GTP, 1.25 mM cap analog 2 and 5 µg/µl plasmid hRLuc-pRNA2(A)128 digested with AarI as a template. Following 2 h incubation, 1 U/µl DNase I was added and incubation was continued for 30 min at 37°C, after which EDTA was added to 25 µM final concentration. Obtained transcripts RNA-1a, RNA-2/3, and RNA-4/5 and also 2-RNARluc were purified using RNA Clean & Concentrator-25 (Zymo Research), NucAway Spin Columns (Ambion), and NucleoSpin RNA Clean-up XS (Macherey-Nagel), respectively. The quality of RNA-1a/1, RNA-2/3/4/5 was checked on 15% acrylamide/7 M



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 20 of 35

urea 1 × TBE gels, while the quality of 2-RNARluc was checked on native 1.2% 1 × TBE agarose gels. In turn, concentrations of all obtained transcripts were determined spectrophotometrically. Translation efficiency in RRL. Following transcription, translation reactions in RRL were performed under conditions determined for capdependent translation, as described previously.(53) A typical reaction mixture (10 µl) contained: 40% RRL lysate (Promega), 0.01 mM mixture of amino acids, 0.6 mM MgCl2, 210 mM potassium acetate, and 5′-capped RNA-3 encoding the firefly luciferase gene; this mixture was incubated at 37°C for 1 h. Three different concentrations (0.1 ng/µl, 0.2 ng/µl, 0.4 ng/µl) of each analyzed transcript were tested by this in vitro translation reaction. The activity of synthesized luciferase was measured using Luciferase Assay System (Promega) in a Glomax Luminometer (Promega), and depicted as a function of the capped luciferase mRNA concentration. After linear fitting to the obtained data points (OriginPro8 software), the translation efficiency of tested transcripts was determined relative to that of the m7GpppG-capped luciferase mRNA (the slope of the fitted line normalized to 1). Each translation reaction mixture was assayed for luciferase activity in duplicate. Post-transcriptional labeling of RNA. Short transcripts. Labeling reactions on short RNAs capped with compound 6 (cap-6-RNA-1a) were performed in borate buffer pH 8.5 at 4.5 µM (51.3 ng/µl) RNA concentration. A stock solution of 5(6)-carboxyfluorescein NHS ester (FAM-NHS) at 0.4 M concentration was either prepared in situ as described above in section “Labeling of cap analogs” (in situ prepared FAM-NHS) or by dissolving 3.79 mg of commercially available fluorescein NHS (Lumiprobe Cat. No 45120) (commercial FAM-NHS) in 20 µl of DMSO. The diluted FAMNHS solutions of 1.35 mM, 13.5 mM, or 135 mM concentrations were prepared by serial dilutions of the stock solution in DMSO. Aliquots (1.2 µl) of each of these solutions were added to the RNA solution (4.8 µl) to achieve 10-, 100- and 1000-fold molar excess of the NHS-ester over RNA, respectively, and 20% final DMSO content. The reaction mixtures were incubated at room temperature on a laboratory shaker for the indicated times (1–6 h). The fluorescently labeled short RNAs were precipitated with ethanol in the presence of 0.3 M sodium acetate and 2 µg of glycogen (Roche) and re-dissolved in deionized water. The RNAs were then trimmed at the 3′ ends with DNAzyme 10-23 (as described in the “RNA synthesis” section, above) and analyzed on 15% acrylamide/7 M urea 1 × TBE gels and visualized using a Typhoon FLA 9500 laser scanner (GE Healthcare) first in FAM-fluorescence mode, then stained with SYBR Gold (Invitrogen). Optimal labeling was achieved within 1 h using 100-fold molar excess of the NHS-ester over RNA. Moreover, the highest labeling efficiency was observed using freshly prepared borate buffer and fluorescein stock solutions. Long transcripts. The labeling reactions for long 6-capped RNAs (6-RNA-4) were performed in borate buffer pH 8.5 at 100 ng/µl RNA concentration using the same protocol as for short RNA labeling, with the exception that a 9.3 mM FAM-NHS solution was used in the labeling reactions to achieve 10000-fold molar excess of NHS ester over RNA. Reactions were incubated for 1 h at room temperature, followed by mRNA purification using NucleoSpin RNA Clean-up XS or NucAway Spin Columns (Ambion) and analyzed on native 1.2% 1 × TBE agarose gels and visualized using a Typhoon FLA 9500 as above, first in FAM-fluorescence mode, then stained with ethidium bromide.


Page 21 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Electrophoretic mobility shift assay (EMSA). For EMSA analysis RNA-2 capped with 5 and 5B was obtained, then 5-RNA-2 transcript was used for posttranscriptional labeling with biotin moieties. Post-transcriptional biotin labeling was performed according to established protocols for fluorescent labeling; 5-RNA-2 transcript was incubated with 10000-fold molar excess of biotin NHS ester over RNA for 1 h at room temperature. Subsequently, both co- and post-transcriptional labeling protocols were tested for interaction with streptavidin. In this analysis, as a negative control, RNA capped with unlabeled analog (5-RNA-2) was used. After denaturation (8 min at 100°C) of biotin-labeled RNAs, they were incubated for 20 minutes on ice with increasing amounts (0–100 pmol, 1 µl) of recombinant tetrameric streptavidin (Promega) in deionized water in a final volume of 6 µl, then 2 µl of ice-cooled sterile 80% glycerol was added. Samples were analyzed on native 1.4% 0.5 × TBE agarose gels, stained with ethidium bromide and visualized under UV illumination using a Gel Doc XR+ (BioRad) gel documentation system. In vitro RNA decapping assay with SpDcp1-Dcp2.

In vitro decapping activity was measured for variously capped RNAs, either short transcripts (RNA-1) or firefly luciferase-encoding mRNAs (RNA-4), as substrates. 15 ng of capped short RNAs or 650 ng of mRNAs were subjected to digestion with 400 nM or 130 nM SpDcp1-Dcp2, respectively in decapping buffer (50 mM Tris-HCl pH 8.0, 50 mM NH4Cl, 0.01% NP-40, 1 mM DTT, and 5 mM MgCl2), in final reaction volumes of 7 µl. Reactions were performed at 37°C for 1 h and terminated by adding an equal volume of loading dye (5 M urea, 44% formamide, 20 mM EDTA, 0.03% bromophenol blue, 0.03% xylene cyanol) for samples containing short RNAs or by adding phenol/chloroform solution to reactions with mRNAs. After phenol/chloroform extraction mRNAs were ethanol precipitated and equal amounts of each samples were loaded on native 1.2% 1 × TBE agarose gel, whereas short RNAs after SpDcp1-Dcp2 treatment were resolved electrophoretically on denaturing 15% acrylamide/7 M urea 1 × TBE gels. Both short and long transcripts were visualized, first in FAMfluorescence mode, then stained with SYBR Gold (Invitrogen) or with ethidium bromide and visualized using a Typhoon FLA 9500 laser scanner (GE Healthcare) or a gel documentation system (Gel Doc XR+, BioRad). Xrn1 treatment. 5′-capped and labeled mRNAs (RNA-4) prior- and post-SpDcp1-Dcp2 treatment were subjected to Xrn1 degradation. A typical reaction mixture (7 µl) contained 70 ng of mRNAs, 0.2 U Xrn1 (New England BioLabs) and 1U/µl RiboLock RNase Inhibitor in NEB Buffer 3 (New England BioLabs). Reactions were performed at 37°C for 1 h and terminated by adding an equal volume of loading dye (95% formamide, 0.5 mM EDTA, 0.025% SDS, 0.03% bromophenol blue, 0.03% xylene cyanol). Reaction products were resolved on native 1.2% 1 × TBE agarose gels and visualized first in FAM-fluorescence mode using a Typhoon FLA 9500, then stained with ethidium bromide and visualized under UV illumination using a gel documentation system, as described above. In vitro RNA decapping assay with hNudt16.

Decapping reactions with hNudt16 enzyme, which was expressed in Escherichia coli and purified as described previously,(54) were performed in 7 µl reaction volumes containing: 100 ng of capped RNA-3 transcripts and 2 µM hNudt16 in reaction buffer (40 mM Tris-HCl pH 7.9, 6 mM MgCl2, 10 mM NaCl, 10 mM DTT, 2 mM



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 22 of 35

spermidine). Reactions were incubated at 30°C for the indicated times and terminated by adding an equal amount of FORMAzol (Molecular Research Center, Inc.) and then incubated for 3 min at 55°C. Reaction products were resolved on native 1% 0.5 × TBE agarose gels and visualized first in FAM-fluorescence mode using a Typhoon FLA 9500, stained with ethidium bromide, and visualized under UV light using a gel documentation system Gel Doc XR+. In vitro degradation assay in HeLa cytoplasmic extracts.

Co- and post-transcriptionally labeled RNA-1a transcripts were subjected to degradation by HeLa cytoplasmic extracts. Human cervical carcinoma HeLa cells were grown in DMEM (Gibco) (with 2 mM l-glutamine) supplemented with 10% FBS (Sigma-Aldrich) and 1% penicillin/streptomycin (Gibco) at 5% CO2 and 37°C. Cytoplasmic extract was prepared as described by Mukherjee and colleagues.(55) Briefly, HeLa cells were washed twice using PBS, than scraped into PBS and centrifuged. The pellet was washed with PBS and with Buffer A (10 mM HEPES pH 7.9, 1.5 mM MgCl2, 10 mM KCl, 1 mM DTT, 1 mM PMSF), and then resuspended in 3 PCV (pellet cell volumes) of Buffer A and incubated on ice for 10 min. Then, the cells were disrupted with a Dounce homogenizer and the lysate was centrifuged at 2000 g for 15 min at 4°C. After centrifugation, supernatant was collected and considered as the cytoplasmic fraction. Then, 0.11 volume of Buffer B (0.3 M HEPES pH 7.9, 30 mM MgCl2, 1.4 M KCl) was added to the cytoplasmic fraction. Prepared extract was centrifuged at 16000 g for 30 min at 4°C and the supernatant supplemented with 10% glycerol and dialyzed against Buffer D (20 mM HEPES pH 7.9, 20% (v/v) glycerol, 100 mM KCl, 0.2 M EDTA, 1 mM DTT) for 2 h, aliquoted and snap-frozen in liquid nitrogen. In vitro enzymatic assays were performed in 25 µl reaction volumes containing 7.5 µl HeLa cytoplasmic extract, 18 mM phosphocreatine, 1.08 mM ATP, 0.4% polyvinyl alcohol, and 30 ng of fluorescently labeled RNA-1a transcripts. Reactions were performed at 30°C for the indicated times and terminated by adding an equal volume of loading dye (5 M urea, 44% formamide, 20 mM EDTA, 0.03% xylene cyanol). Reaction products were resolved on denaturing 15% acrylamide/7 M urea 1 × TBE gels and visualized with Typhoon FLA 9500 using a FAM-fluorescence mode. mRNA visualization in HeLa cells. HeLa cells were cultured as described above (“In vitro degradation assay in HeLa cytoplasmic extracts”). In a typical experiment, 24 h before transfection, 5 × 104 cells were seeded per well with glass coverslips in 24-well plates. Before transfection, cells were washed with PBS and the medium was changed to Opti-MEM (Gibco). Cells were transfected using 1.5 µl Lipofectamine MessengerMAX Transfection Reagent (Invitrogen) and 0.5 µg mRNA (RNA-5). Following transfection for 5 h at 37°C cells were washed with PBS and fixed using 4% paraformaldehyde in PBS for 15 min at room temperature. Then, the nuclei were stained using Hoechst (SigmaAldrich). Cell samples were imaged using a Carl Zeiss LSM 700 confocal laser scanning microscope with Axio Imager Z2, using a 40 × /1.3 oil objective lens. Hoechst and fluorescein emissions were detected at 300–493 nm and 493–800 nm, respectively, after excitation at 405 nm for Hoechst and 488 nm for fluorescein. mRNA translation in HeLa cells. HeLa cells were cultured as described above (“In vitro degradation assay in HeLa cytoplasmic extracts”). In a typical experiment, 24 h before transfection, 104 cells per well were seeded in a 96-well plate. Directly before


Page 23 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


transfection, cells were washed with PBS and the medium was changed to Opti-MEM. Cells were transfected using 0.3 µl Lipofectamine MessengerMAX Transfection Reagent and 70 ng mRNA encoding firefly luciferase (RNA-5) and 30 ng control mRNA encoding Renilla luciferase and capped with compound 2 (2-RNARluc). Measurement for each pair of transcripts was performed in triplicate. Following transfection for 5 h at 37°C cells were washed with PBS and lysed using the Dual-Luciferase Reporter Assay System (Promega) followed by determination of luciferase activity. Bioluminescent flux was measured with the Synergy H1 microplate reader (BioTek). Data were normalized in order to account for differences in transfection efficiencies in particular experiments. Normalization was performed according to the equation: Fnorm = F/R·R2-RNARluc_AVG, where, F = firefly luminescence readout for a chosen well, R = Renilla luminescence readout for the same well as F, R2RNARluc_AVG

= average Renilla luminescent value for all transfections, Fnorm = normalized luciferase activity value

(data shown in Fig. 9). ACKNOWLEDGEMENTS The authors are grateful to Stephen R. Ikeda (The National Institute on Alcohol Abuse and Alcoholism) for providing hRLuc-pRNA2(A)128 plasmid and John D. Gross (University of California, San Francisco) for providing plasmid encoding Dcp1-Dcp2 complex from Schizosaccharomyces pombe. ASSOCIATED CONTENT The Supporting Information is available: Table S1, Figures S1–S4, Chemical synthesis procedures and products characterization including HPLC profiles, ESI(-) HR MS spectra, UV-Vis and fluorescence spectra. FUNDING SOURCES This work was supported by the National Science Centre, Poland (grant no UMO-2013/09/B/ST5/01341 to JJ); the National Science Centre, Poland (grant no UMO-2013/08/A/NZ1/00866 to ED), the Ministry of Science and Higher Education, Poland (Grant No. DI2012 024842 to MW), the Polish National Center of Research and Development (STRATEGMED1/235773/19/NCBR/2016 to JJ and ED). REFERENCES

(1) (2) (3)

(4) (5) (6)

Kushner, S. R. (2002) mRNA Decay in Escherichia coli Comes of Age. J. Bacteriol. 184, 46584665. Yu, J., and Russell, J. E. (2001) Structural and Functional Analysis of an mRNP Complex That Mediates the High Stability of Human β-Globin mRNA. Mol. Cell. Biol. 21, 5879-5888. Kempe, H., Schwabe, A., Crémazy, F., Verschure, P. J., and Bruggeman, F. J. (2015) The volumes and transcript counts of single cells reveal concentration homeostasis and capture biological noise. Mol. Biol. Cell 26, 797-804. Weil, T. T., Parton, R. M., and Davis, I. (2010) Making the message clear: visualizing mRNA localization. Trends Cell Biol. 20, 380-390. Rombouts, K., Braeckmans, K., and Remaut, K. (2016) Fluorescent Labeling of Plasmid DNA and mRNA: Gains and Losses of Current Labeling Strategies. Bioconjugate Chem. 27, 280-297. Paredes, E., and Das, S. R. (2011) Click Chemistry for Rapid Labeling and Ligation of RNA. ChemBioChem 12, 125-131. 23 ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

(7)

(8)

(9)

(10) (11)

(12)

(13)

(14)

(15) (16)

(17) (18) (19)

(20) (21) (22)

(23)

(24)

(25)

Page 24 of 35

Fauster, K., Hartl, M., Santner, T., Aigner, M., Kreutz, C., Bister, K., Ennifar, E., and Micura, R. (2012) 2′-Azido RNA, a Versatile Tool for Chemical Biology: Synthesis, X-ray Structure, siRNA Applications, Click Labeling. ACS Chem. Biol. 7, 581-589. Sawant, A. A., Tanpure, A. A., Mukherjee, P. P., Athavale, S., Kelkar, A., Galande, S., and Srivatsan, S. G. (2016) A versatile toolbox for posttranscriptional chemical labeling and imaging of RNA. Nucleic Acids Res. 44, e16-e16. Li, Y., Fin, A., McCoy, L., and Tor, Y. (2017) Polymerase-Mediated Site-Specific Incorporation of a Synthetic Fluorescent Isomorphic G Surrogate into RNA. Angew. Chem.-Int. Edit. 56, 1303-1307. Jao, C. Y., and Salic, A. (2008) Exploring RNA transcription and turnover in vivo by using click chemistry. Proc. Natl. Acad. Sci. U. S. A. 105, 15779-15784. Rovira, A. R., Fin, A., and Tor, Y. (2017) Expanding a fluorescent RNA alphabet: synthesis, photophysics and utility of isothiazole-derived purine nucleoside surrogates. Chemical Science 8, 2983-2993. Lavergne, T., Lamichhane, R., Malyshev, D. A., Li, Z., Li, L., Sperling, E., Williamson, J. R., Millar, D. P., and Romesberg, F. E. (2016) FRET Characterization of Complex Conformational Changes in a Large 16S Ribosomal RNA Fragment Site-Specifically Labeled Using Unnatural Base Pairs. ACS Chem. Biol. 11, 1347-1353. Seo, Y. J., Malyshev, D. A., Lavergne, T., Ordoukhanian, P., and Romesberg, F. E. (2011) SiteSpecific Labeling of DNA and RNA Using an Efficiently Replicated and Transcribed Class of Unnatural Base Pairs. J. Am. Chem. Soc. 133, 19878-19888. Kawai, R., Kimoto, M., Ikeda, S., Mitsui, T., Endo, M., Yokoyama, S., and Hirao, I. (2005) SiteSpecific Fluorescent Labeling of RNA Molecules by Specific Transcription Using Unnatural Base Pairs. J. Am. Chem. Soc. 127, 17286-17295. Pyka, A. M., Domnick, C., Braun, F., and Kath-Schorr, S. (2014) Diels–Alder Cycloadditions on Synthetic RNA in Mammalian Cells. Bioconjugate Chem. 25, 1438-1443. Domnick, C., Eggert, F., and Kath-Schorr, S. (2015) Site-specific enzymatic introduction of a norbornene modified unnatural base into RNA and application in post-transcriptional labeling. Chem. Comm. 51, 8253-8256. Eggert, F., and Kath-Schorr, S. (2016) A cyclopropene-modified nucleotide for site-specific RNA labeling using genetic alphabet expansion transcription. Chem. Comm. 52, 7284-7287. Santangelo, P. J., Nix, B., Tsourkas, A., and Bao, G. (2004) Dual FRET molecular beacons for mRNA detection in living cells. Nucleic Acids Res. 32, e57-e57. Briley, W. E., Bondy, M. H., Randeria, P. S., Dupper, T. J., and Mirkin, C. A. (2015) Quantification and real-time tracking of RNA in live cells using Sticky-flares. Proc. Natl. Acad. Sci. U. S. A. 112, 9591-9595. Paige, J. S., Wu, K. Y., and Jaffrey, S. R. (2011) RNA Mimics of Green Fluorescent Protein. Science 333, 642. Sparano, B. A., and Koide, K. (2007) Fluorescent Sensors for Specific RNA: A General Paradigm Using Chemistry and Combinatorial Biology. J. Am. Chem. Soc. 129, 4785-4794. McDonald, R. I., Guilinger, J. P., Mukherji, S., Curtis, E. A., Lee, W. I., and Liu, D. R. (2014) Electrophilic activity-based RNA probes reveal a self-alkylating RNA for RNA labeling. Nat. Chem. Biol. 10, 1049-1054. Sharma, A. K., Plant, J. J., Rangel, A. E., Meek, K. N., Anamisis, A. J., Hollien, J., and Heemstra, J. M. (2014) Fluorescent RNA Labeling Using Self-Alkylating Ribozymes. ACS Chem. Biol. 9, 1680-1684. Jahn, K., Olsen, E. M., Nielsen, M. M., Tørring, T., MohammadZadegan, R., Andersen, E. S., Gothelf, K. V., and Kjems, J. (2011) Site-Specific Chemical Labeling of Long RNA Molecules. Bioconjugate Chem. 22, 95-100. Li, F., Dong, J., Hu, X., Gong, W., Li, J., Shen, J., Tian, H., and Wang, J. (2015) A Covalent Approach for Site-Specific RNA Labeling in Mammalian Cells. Angew. Chem.-Int. Edit. 54, 4597-4602. 24 ACS Paragon Plus Environment

Page 25 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(26)

(27)

(28)

(29)

(30) (31) (32)

(33)

(34) (35)

(36)

(37) (38) (39)

(40)

(41)

(42)

(43)

Winz, M.-L., Samanta, A., Benzinger, D., and Jäschke, A. (2012) Site-specific terminal and internal labeling of RNA by poly(A) polymerase tailing and copper-catalyzed or copper-free strain-promoted click chemistry. Nucleic Acids Res. 40, e78-e78. Alexander, S. C., Busby, K. N., Cole, C. M., Zhou, C. Y., and Devaraj, N. K. (2015) Site-Specific Covalent Labeling of RNA by Enzymatic Transglycosylation. J. Am. Chem. Soc. 137, 1275612759. Muttach, F., and Rentmeister, A. (2016) A Biocatalytic Cascade for Versatile One-Pot Modification of mRNA Starting from Methionine Analogues. Angew. Chem.-Int. Edit. 55, 1917-1920. Holstein, J. M., Muttach, F., Schiefelbein, S. H. H., and Rentmeister, A. (2017) Dual 5′ Cap Labeling Based on Regioselective RNA Methyltransferases and Bioorthogonal Reactions. Chem. - Eur. J. 23, 6165-6173. Schulz, D., Holstein, J. M., and Rentmeister, A. (2013) A Chemo-Enzymatic Approach for SiteSpecific Modification of the RNA Cap. Angew. Chem.-Int. Edit. 52, 7874-7878. Holstein, J. M., Schulz, D., and Rentmeister, A. (2014) Bioorthogonal site-specific labeling of the 5[prime or minute]-cap structure in eukaryotic mRNAs. Chem. Comm. 50, 4478-4481. Holstein, J. M., Stummer, D., and Rentmeister, A. (2015) Enzymatic modification of 5[prime or minute]-capped RNA with a 4-vinylbenzyl group provides a platform for photoclick and inverse electron-demand Diels-Alder reaction. Chemical Science 6, 1362-1369. Holstein, J. M., Anhäuser, L., and Rentmeister, A. (2016) Modifying the 5′-Cap for Click Reactions of Eukaryotic mRNA and To Tune Translation Efficiency in Living Cells. Angew. Chem.-Int. Edit. 55, 10899-10903. Ogasawara, S. (2017) Duration Control of Protein Expression in Vivo by Light-Mediated Reversible Activation of Translation. ACS Chem. Biol. 12, 351-356. Jemielity, J., Lukaszewicz, M., Kowalska, J., Czarnecki, J., Zuberek, J., and Darzynkiewicz, E. (2012) Synthesis of biotin labelled cap analogue - incorporable into mRNA transcripts and promoting cap-dependent translation. Org. Biomol. Chem. 10, 8570-8574. Ziemniak, M., Szabelski, M., Lukaszewicz, M., Nowicka, A., Darzynkiewicz, E., Rhoads, R. E., Wieczorek, Z., and Jemielity, J. (2013) Synthesis and evaluation of fluorescent cap analogues for mRNA labelling. RSC Adv. 3, 20943-20958. Ogasawara, S., and Maeda, M. (2011) Photoresponsive 5 ′ -cap for the reversible photoregulation of gene expression. Bioorg. Med. Chem Lett. 21, 5457-5459. Hiremath, L. S., Webb, N. R., and Rhoads, R. E. (1985) Immunological detection of the messenger RNA cap-binding protein. J. Biol. Chem. 260, 7843-7849. Duncan, R., Milburn, S. C., and Hershey, J. W. (1987) Regulated phosphorylation and low abundance of HeLa cell initiation factor eIF-4F suggest a role in translational control. Heat shock effects on eIF-4F. J. Biol. Chem. 262, 380-388. Jemielity, J., Kowalska, J., Rydzik, A. M., and Darzynkiewicz, E. (2010) Synthetic mRNA cap analogs with a modified triphosphate bridge - synthesis, applications and prospects. New J. Chem. 34, 829-844. Grudzien-Nogalska, E., Kowalska, J., Su, W., Kuhn, A. N., Slepenkov, S. V., Darzynkiewicz, E., Sahin, U., Jemielity, J., and Rhoads, R. E. (2013) Synthetic mRNAs with Superior Translation and Stability Properties, in Synthetic Messenger RNA and Cell Metabolism Modulation: Methods and Protocols (Rabinovich, P. M., Ed.) pp 55-72, Humana Press, Totowa, NJ. Warminski, M., Warminska, Z., Kowalska, J., and Jemielity, J. (2015) mRNA Cap Modification through Carb-amate Chemistry: Synthesis of Amino- and Carboxy-Functionalised Cap Analogues Suitable for Labelling and Bioconjugation. Eur. J. Org. Chem. 2015, 6153-6169. Song, M.-G., Bail, S., and Kiledjian, M. (2013) Multiple Nudix family proteins possess mRNA decapping activity. RNA 19, 390-399.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

(44)

(45)

(46) (47)

(48) (49)

(50)

(51)

(52)

(53)

(54)

(55)

Page 26 of 35

Lu, G., Zhang, J., Li, Y., Li, Z., Zhang, N., Xu, X., Wang, T., Guan, Z., Gao, G. F., and Yan, J. (2011) hNUDT16: a universal decapping enzyme for small nucleolar RNA and cytoplasmic mRNA. Protein Cell 2, 64-73. Melton, D. A., Krieg, P. A., Rebagliati, M. R., Maniatis, T., Zinn, K., and Green, M. R. (1984) Efficient in vitro synthesis of biologically active RNA and RNA hybridization probes from plasmids containing a bacteriophage SP6 promoter. Nucleic Acids Res. 12, 7035-7056. Yisraeli, J. K., and Melton, D. A. (1989) [4] Synthesis of long, capped transcripts in Vitro by SP6 and T7 RNA polymerases, in Methods in Enzymology pp 42-50, Academic Press. Zuberek, J., Wyslouch-Cieszynska, A., Niedzwiecka, A., Dadlez, M., Stepinski, J., Augustyniak, W., Gingras, A.-C., Zhang, Z., Burley, S. K., Sonenberg, N., et al. (2003) Phosphorylation of eIF4E attenuates its interaction with mRNA 5′ cap analogs by electrostatic repulsion: Inteinmediated protein ligation strategy to obtain phosphorylated protein. RNA 9, 52-61. Floor, S. N., Jones, B. N., Hernandez, G. A., and Gross, J. D. (2010) A split active site couples cap recognition by Dcp2 to activation. Nat. Struct. Mol. Biol. 17, 1096-1101. Niedzwiecka, A., Marcotrigiano, J., Stepinski, J., Jankowska-Anyszka, M., Wyslouch-Cieszynska, A., Dadlez, M., Gingras, A.-C., Mak, P., Darzynkiewicz, E., Sonenberg, et al. (2002) Biophysical Studies of eIF4E Cap-binding Protein: Recognition of mRNA 5′ Cap Structure and Synthetic Fragments of eIF4G and 4E-BP1 Proteins. J. Mol. Biol. 319, 615-635. Strenkowska, M., Kowalska, J., Lukaszewicz, M., Zuberek, J., Su, W., Rhoads, R. E., Darzynkiewicz, E., and Jemielity, J. (2010) Towards mRNA with superior translational activity: synthesis and properties of ARCA tetraphosphates with single phosphorothioate modifications. New J. Chem. 34, 993-1007. Kowalska, J., Lukaszewicz, M., Zuberek, J., Darzynkiewicz, E., and Jemielity, J. (2009) Phosphoroselenoate Dinucleotides for Modification of mRNA 5′ End. ChemBioChem 10, 2469-2473. Williams, D., Puhl, H., and Ikeda, S. (2010) A Simple, Highly Efficient Method for Heterologous Expression in Mammalian Primary Neurons Using Cationic Lipid-mediated mRNA Transfection. Front. Neurosci. 4, 181. Rydzik, A. M., Lukaszewicz, M., Zuberek, J., Kowalska, J., Darzynkiewicz, Z. M., Darzynkiewicz, E., and Jemielity, J. (2009) Synthetic dinucleotide mRNA cap analogs with tetraphosphate 5',5' bridge containing methylenebis(phosphonate) modification. Org. Biomol. Chem. 7, 4763-4776. Wojtczak, B. A., Warminski, M., Kowalska, J., Lukaszewicz, M., Honcharenko, M., Smith, C. I. E., Stromberg, R., Darzynkiewicz, E., and Jemielity, J. (2016) Clickable trimethylguanosine cap analogs modified within the triphosphate bridge: synthesis, conjugation to RNA and susceptibility to degradation. RSC Adv. 6, 8317-8328. Mukherjee, D., Fritz, D. T., Kilpatrick, W. J., Gao, M., and Wilusz, J. (2004) Analysis of RNA Exonucleolytic Activities in Cellular Extracts, in mRNA Processing and Metabolism: Methods and Protocols (Schoenberg, D. R., Ed.) pp 193-211, Humana Press, Totowa, NJ.


Page 27 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Structure of eukaryotic mRNA 5′ end (m7G cap). 84x56mm (300 x 300 DPI)

ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Structures and nomenclature of mRNA 5′ cap analogs and their conjugates studied in this work. A) Reference analogs (1-3): m7GpppG (1) – standard m7G cap analog; m27,3′-O GpppG (2) – anti-reverse cap analog or ARCA, yields mRNAs with translation efficiency superior to 1; ApppG (3) – non-functional cap, yields mRNAs with poor translation efficiency. B) Cap analogs 4-6 carrying NH2 groups to enable mRNA 5′ end labeling: cap with short linker (4), cap with medium linker (5), cap with long linker (6). Label attachment site is marked with Z (for unlabeled analog Z = H). All analogs were synthesized as a mixture of 2′-O and 3′-O regioisomers (ratio of 2′-O:3′-O isomers were as follows: 1:1.79 for 4, 1:1.16 for 5, 1:1.24 for 6); C) Labels (Z) used in experiments: biotin (B), fluorescein (F), and acetyl (Ac). Linkers are colored blue; labels are colored red. 163x124mm (300 x 300 DPI)


Page 28 of 35

Page 29 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Biochemical properties of 2′-O and 3′-O substituted cap analogs. A) Association constants (KAS) for complexes of translation initiation factor 4E (eIF4E) and selected 5′ cap analogs (isolated 2′-O and 3′-O isomers, a and b, respectively). Data shown are means from triplicate experiments. B) Relative translation efficiencies of luciferase mRNAs capped with selected analogs (2′-O and 3′-O mixtures or isolated isomers) in rabbit reticulocyte lysate (RRL). Results are color coded according to the type of modification present. Data shown represent means from duplicate experiments. 159x73mm (300 x 300 DPI)



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

A) Two approaches to mRNA 5′ cap labeling explored in this work: co-transcriptional (co-trx) and posttranscriptional (post-trx) labeling; B) NHS-labeling of amino-functionalized cap analog; Label: 5(6)carboxyfluorescein, biotin or acetic acid; TSTU: N,N,N’,N’-Tetramethyl-O-(N-succinimidyl)uronium tetrafluoroborate 213x64mm (300 x 300 DPI)


Page 30 of 35

Page 31 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Demonstration of post-transcriptional labeling procedures for RNAs of various lengths. A) Electrophoretic analysis of post-transcriptionally FAM-labeled 24-nt RNA-1 capped with cap 6a (labeling conditions: 1 h, 100-fold excess of FAM-NHS obtained by in situ activation). Uncapped and m7GpppG-capped RNAs were used as negative controls. B) Electrophoretic analysis of post-transcriptionally FAM-labeled luciferase mRNAs (RNA-4) capped with cap 6 (labeling conditions: 1 h, 10,000-fold excess of FAM-NHS obtained either by in situ activation or from a commercial source). Uncapped RNA and RNAs co-transcriptionally capped with m7GpppG and cap 5F were used as controls. Whole gel images are shown in the Supporting Information (Figure S4). C) Electrophoretic mobility shift assay showing formation of the complexes of streptavidin and 140-nt RNAs (RNA-2) biotinylated within 5′ cap structures by either the co-transcriptional approach (transcription reaction in the presence of cap 5B) or the post-transcriptional approach (reaction of 5B-RNA-2 with NHS-biotin). 1-RNA-2 was used as a negative control. 212x77mm (300 x 300 DPI)



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Utility of fluorescently labeled transcripts in RNA decapping assays. A) Dcp2/Dcp1-catalyzed decapping of RNA-1 (24 nt) obtained by either a post- or co-transcriptional protocol. B) Electrophoretic analysis of posttranscriptionally labeled luciferase mRNAs before and after incubation with decapping enzyme SpDcp1-Dcp2 followed by incubation with Xrn1. Gels were visualized both in fluorescence mode and by SYBR® Gold or EtBr staining; mRNAs co-transcriptionally labeled with 5F were used as a control. Whole gel images are shown in Supporting information (Figure S6). 239x69mm (300 x 300 DPI)


Page 32 of 35

Page 33 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Utility of fluorescently labeled transcripts in monitoring of 3′-to-5′ RNA decay. Electrophoretic analysis of RNA-1a (35 nt) labeled by a post- (A) and co-transcriptional (B) protocol incubated with HeLa cytoplasmic extract for the indicated times. Heat-denatured extract (–) was used as a negative control. Lanes with fluorescently labeled cap (6aF, 5F) are shown for reference. 169x51mm (300 x 300 DPI)



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Localization of fluorescently labeled mRNA in HeLa cells. HeLa cells were transfected with cotranscriptionally (5F-RNA-5 co-trx) or post-transcriptionally (6aF-RNA-5 post-trx) labeled mRNA encoding firefly luciferase. HeLa cells transfected with mRNA capped with unlabeled analog 6a (6a-RNA-5) and mocktreated cells were used as negative controls. Panels from left to right show fluorescein fluorescence, Hoechst fluorescence, merge of these two panels (merge 1), bright field and merge of all images. Scale bars are 20 µm. 144x114mm (300 x 300 DPI)


Page 34 of 35

Page 35 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Translation efficiencies in HeLa cells for differently capped firefly luciferase mRNAs (RNAs-5). Cells were transfected for 5 h with a mixture of 70 ng luciferase mRNA (RNA-5), uncapped or carrying the studied cap structure, and 30 ng control Renilla luciferase mRNA capped with analog 2 (2-RNARluc). The activity of both luciferases was measured using a dual-reporter assay, followed by normalization of firefly luciferase activity to Renilla activity to account for transfection efficiency differences as described in the experimental section. “2”, “3”, “6a”, “5F” denote RNA-5 transcripts co-transcriptionally capped at their 5′ ends with cap analogs 2, 3, 6a, 5F, respectively. “pppG” denotes uncapped (GTP-initiated) RNA-5, while “2+F” and “6a+F” refer to RNA-5 capped with analogs 2 and 6a, respectively, subjected to post-transcriptional labeling procedure with commercial FAM-NHS. “Mock” refers to cells transfected only with control RNA (2-RNARluc). Data shown are average values from triplicate experiments ± S.D. Values above each bar represent mean normalized luciferase activity. 224x179mm (300 x 300 DPI)


Amino-functionalized 5' cap analogs as tools for site-specific

Recommend Documents