This work was supported by grants from the Wellcome Trust (091549 to S.G and 109093/Z/15/A to S.M.), the Wellcome Trust Centre for Cell Biology core grant (092076) and Medical Research Council Non-Clinical Senior Research Fellowship (MR/R008205/1 to S.G.), the European Molecular Biology Organization under a long-term postdoctoral fellowship (ALTF 1070-2017 to R.A.C), and the Independent Research Fund Denmark (T.H.J).

A. Langford and W. Worboys are affiliated with UVO3, a commercial company. They had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

The &#967;CRAC method, combined with the new cross-linking and cell harvesting devices, has great potential because it is applicable to a wide range of model organisms and therefore should be of general interest to the RNA field. There are many areas in which &#967;CRAC can be utilized. For example, the method could be used to measure the hierarchical assembly of proteins into large macromolecular complexes, such as the spliceosome and the ribosome, which often involves dynamic interactions between proteins and RNA molecules. We also now routinely use it to monitor interactions between RNA decay factors and their substrates when cells are subjected to diverse kind of stresses. This enables us to determine at what stage of the adaptive response these factors are most active, what substrates they bind to, and how dynamic these interactions are. Such data should enable researchers to determine the relative contribution of each factor in adaptation to environmental changes.
&#967;CRAC uses dual affinity purification tags (HTF or HTP) to purify the protein under highly stringent and denaturing conditions. This ensures that the copurified RNA is highly enriched for RNAs that were covalently cross-linked to the protein of interest. However, relying on affinity tags has disadvantages. For example, the tag could interfere with protein function, which could give a distorted readout of its RNA-binding interactome. Additionally, for some model organisms it may not always be possible to utilize tags because the genetic tools to integrate DNA fragments into the genome or to transform expression plasmids are not yet available. However, it is straightforward to alter some parts of the &#967;CRAC protocol to make it compatible with CLIP-based protocols that rely on antibodies for purification of the RBP. Indeed, this study showed that it is possible to combine iCLIP-based purifications with our crosslinker. We are now in the process of developing CLIP protocols to study the temporal association of human RNA-binding proteins with nascent RNA transcripts.
When performing &#967;CRAC on a new protein, the UV exposure must be optimized in order to induce maximal cross-linking. This is important&#160;because high UV exposures can reduce the recovery of RNA during the purification step. Cells expressing the recombinant RBP were exposed to various UV doses, 100 mJ/cm2, 250 mJ/cm2, 500 mJ/cm2, and 1 J/cm2. The RNPs were then captured and the RNAs were fragmented and radiolabeled. Afterwards, the RNPs were resolved by SDS-PAGE and an autoradiogram was taken in order to deduce which exposure gave the most intense signal (i.e. the maximal cross-linking).
Once the experimental conditions are optimized, several control experiments are recommended when performing &#967;CRAC. First, a UV irradiated, untagged sample can be used to monitor background binding to the purification beads. Second, when applying &#967;CRAC during a shift experiment, a second time series where the cells are shifted back into the original medium enables investigation into whether the filtration of the cells itself induces changes in RNA levels or protein-RNA interactions.
As mentioned in the Introduction, numerous recently published papers suggest a number of optimizations to the CLIP protocol. This includes the use of fluorescently labeled adapters for detecting the protein-RNA complex through infrared scanning10 as well as optimizations to various nucleic acid purification and size selection steps shown to increase the complexity of the resulting libraries12,45. We are currently implementing some of these improvements to further refine the &#967;CRAC protocol. The protocol presented here already contains a number of improvements to the original CRAC and &#967;CRAC protocols that increase the complexity of the data. For example, previously, after resolving the cross-linked, radioactive protein-RNA complexes on SDS-PAGE gels, they were transferred to a nitrocellulose membrane and the cross-linked RNA was isolated from the blot. However, the transfer of the RNP and subsequent RNA extraction can be very inefficient, particularly when dealing with large RBPs such as RNA polymerase subunits. This can result in a significant reduction in the recovery of the cross-linked RNA. In the current protocol, the cross-linked RNA is extracted directly from SDS-PAGE gel slices as illustrated in Figure 1. This increased the recovery of cross-linked RNAs. Additionally, after PCR amplification of the cDNAs the product was originally resolved on 3%, low melting temperature agarose gels, and then 175&#8211;300 bp PCR products were extracted from the gel. However, these gels can be easily overloaded, resulting in very poor separation of the DNA. Replacing agarose gels with precast TBE gels resulted in more consistent size separation and better recovery of PCR products.

An erratum was issued for:&#160;<a href="https://www.jove.com/t/61027">Monitoring Protein-RNA Interaction Dynamics in vivo at High Temporal Resolution using &#967;CRAC</a>. The Authors section was updated from:
Stuart W. McKellar1 
Ivayla Ivanova1 
Robert W. van Nues2 
Ross A. Cordiner3 
Will Worboys4 
Andrew Langford4 
Torben Heick Jensen3 
Sander Granneman1 
1Centre for Synthetic and Systems Biology, University of Edinburgh 
2Institute of Cell Biology, University of Edinburgh 
3Department of Molecular Biology and Genetics, Aarhus University, 4UVO3 Ltd.
to:
Stuart W. McKellar1 
Ivayla Ivanova1 
Robert W. van Nues2 
Ross A. Cordiner3 
Mehak Chauhan1 
Niki&#160;Christopoulou1 
Will Worboys4 
Andrew Langford4 
Torben Heick Jensen3 
Sander Granneman1 
1Centre for Engineering Biology, University of Edinburgh 
2Institute of Cell Biology, University of Edinburgh 
3Department of Molecular Biology and Genetics, Aarhus University, 4UVO3 Ltd.

An erratum was issued for:&#160;<a href="https://www.jove.com/t/61027">Monitoring Protein-RNA Interaction Dynamics in vivo at High Temporal Resolution using &#967;CRAC</a>. The Authors section was updated.

Erratum: Monitoring Protein-RNA Interaction Dynamics in vivo at High Temporal Resolution using &#967;CRAC

RNAs often rely on RBPs to exert their function, which has led to great interest in understanding the dynamics between these molecules. Many RBPs have been identified in a wide variety of organisms. However, it has always been notoriously difficult to study RBP-RNA interactions in vivo. A major breakthrough in studying such interactions came with the emergence of CLIP1. This method utilizes ultraviolet (UV, 254 nm) irradiation to induce covalent bonds between RBPs and their directly bound RNAs (i.e., zero-distance cross-linking). Subsequently, the RBP of interest is immunopurified under stringent conditions to ensure that only RNAs covalently cross-linked to the proteins are identified. Bound RNAs are then partially digested with RNases and subsequently converted into cDNA libraries for sequencing. The high purification stringency is important as it greatly increases the specificity of protein and RNA recovery, which is also further enhanced through SDS-PAGE purification of the cross-linked ribonucleoprotein (RNP) complex. CLIP and related methods also provide nucleotide resolution insight into the protein binding site, because during the preparation of the sequencing library, amino acids that cross-linked to the RNA frequently terminate the reverse transcriptase or cause the enzyme to introduce mutations at this site1,2,3.
Since its introduction, the original CLIP protocol has produced a remarkable variety of derivative methodologies. A particularly important breakthrough came with the development of HITS-CLIP (or CLIP-seq), which merges high-throughput sequencing with the CLIP approach3. This has since been adopted by all CLIP-based methodologies. iCLIP introduced improvements in the RNase-mediated trimming and adaptor ligation techniques that facilitate more accurate mapping of the RBP binding sites4. PAR-CLIP combined 4thio-uridine/uracil labeling with cross-linking at 365 nm, making it possible to map cross-linking sites by analyzing T-C substitutions5. CRAC, urea-iCLIP, dCLIP, and uvCLAP introduced denaturing conditions and dual affinity purification steps that further reduce background binding to the affinity resin and further increase the specificity of protein capture2,6,7,8,9. Additionally, CRAC, uvCLAP, and dCLIP introduced tagging the RBP of interest with an affinity tag, thus overcoming the need to generate specific antibodies.
Several optimizations have also been made to expedite the CLIP methodology. The original CLIP protocol utilized radiolabeling of the captured RNAs in order to visualize the RBP-RNA complexes after SDS-PAGE. However, the use of radioactivity can be problematic for laboratories not set up for such work. irCLIP incorporates a fluorophore-coupled adaptor that facilitates visualization through infrared imaging10 and sCLIP utilizes biotinylation of captured RNAs in order to visualize them through streptavidin-conjugated HRP11. Furthermore, eCLIP completely forgoes RNA labeling; instead the protein is excised based solely on its known size12. Streptavidin-based purification has also been used to speed up the process of library preparation in FAST-iCLIP, where a biotinylated 3&#8217; adaptor is ligated onto the RNAs and used to enable purification after reverse transcription and circularization13. Additional enhancements to the iCLIP protocol also greatly increased the complexity of the libraries4.
Finally, CLIP has been modified to enable capture of RBPs from different cellular subcompartments14,15, to visualize newly transcribed RNAs using pulsed induction of photoactivable ribonucleosides5,16,17, to capture methylated RNAs18,19,20, to examine RNA-RNA interactions21,22, and to map 3&#8217; ends23,24.
Despite the great contributions of CLIP-based techniques in aiding our understanding of the interactions between RBPs and RNAs, it has been limited by the inefficiency of UV cross-linking. Although culture cells grown in a monolayer are generally relatively easy to cross-link, this is significantly more challenging in tissues or cells in solution. Tissues can require multiple rounds of UV exposure in order to penetrate to the required cell layers, while microbial cells are often grown in rich mediums that contain aromatic, UV-absorbing compounds25. Indeed, UV irradiation times of up to 30 min have been used to generate sufficient cross-linking between RBPs and their bound RNAs for such samples26,27,28. This extended UV exposure induces stress responses within the cell, such UV-induced DNA damage, which can contaminate the final data in some applications.
The majority of CLIP studies have focused on generating single &#34;snapshots&#34; of specific protein-RNA interactions in a cell. However, protein-RNA interactions are inherently dynamic, particularly when cells are subject to changes in their environment. This can include a sudden reduction in the availability of essential nutrients or rapid changes in temperature. As such, to truly understand the role of an RBP during stress, it is best to perform time-resolved analyses because they can capture the full spectrum of RBP targets during stress and determine at what stage of the stress response the chosen RBP is active. In particular, studies in yeast showed that the first few minutes of adaptation are absolutely crucial for survival and RNA half-lives in bacteria can vary from minutes to seconds29,30,31,32,33. Therefore, such time-resolved analyses should ideally be performed at high temporal resolution. However, the long cross-linking times make the study of early stage adaptive responses particularly challenging.
In order to overcome these issues, we recently developed an improved method that is capable of cross-linking and harvesting cells on minute-long timescales. Our &#967;CRAC method allows quantitative measurement of dynamic changes in RBP-RNA interactions at previously unwitnessed resolution. Crucial to this method was the development of a novel UV irradiation device32 that reduces the required cross-linking time in yeast and bacteria in solution around 10-fold, effectively freezing RBP-RNA interactions instantaneously. In addition, in order to rapidly harvest the cells after UV irradiation, we developed a vacuum filtration device that can harvest exponentially growing yeast in a 0.5 L culture in around 30 s32. These technological innovations allow the study of RBP-RNA dynamics at minute-scale resolution. Additionally, we also introduced several optimizations to the original CRAC protocol2 in order to increase its practicality.
Using &#967;CRAC, we recently studied the targetome of a yeast nuclear RBP, Nab3, in response to glucose deprivation. In Saccharomyces cerevisiae, Nab3 can form a complex with Nrd1, an RBP, and the RNA helicase Sen1 to form the NNS complex. NNS binding to the RNA polymerase and the nascent transcript can trigger transcriptional termination34. This complex is mostly involved in removing cryptic noncoding RNA transcripts but has also been shown to regulate expression of protein-coding genes. The study showed differential targeting of Nab3 to noncoding and coding transcripts after only a minute of stress32. We demonstrated that co-transcriptional termination by Nab3 results in a very transient, pulse-like expression of retrotransposon genes, which would have been difficult to detect using traditional CLIP-based approaches. Additionally, the short UV irradiation times in our UV cross-linker also significantly increased the recovery of short-lived noncoding RNAs32. &#967;CRAC will likely be a crucial tool in elucidating not only how RBPs shape the response to stress on immediate timescales but also their changing roles during the whole lifecycle of a response. This manuscript provides a detailed overview of all the steps in the &#967;CRAC protocol. For illustrative purposes, the method was used to study the yeast Nrd1 protein, which is involved in transcriptional termination and RNA decay35,36, and its RNA targetome in response to glucose deprivation across a multitude of timepoints. Finally, we also demonstrate that our UV irradiation unit can rapidly cross-link RBPs to RNA in HeLa cells, making it possible to also perform high-resolution time-resolved analyses in adherent cells.

<table><tbody><tr><td>1,4-dithioreitol</td><td>Merck</td><td>10708984001</td><td>Buffer component in mammalian cell lysis</td></tr><tr><td>1.5 mL tubes</td><td>Eppendorf</td><td>0030 120.086</td><td>General reaction tube</td></tr><tr><td>2 mL tubes</td><td>Eppendorf</td><td>0030 123.344</td><td>For holding columns and collection of waste</td></tr><tr><td>32P-yATP</td><td>Perkin Elmer</td><td>NEG502Z-250</td><td>For radiolabelling the 5&#039; end of the RNA</td></tr><tr><td>4-12% Bis-Tris gel</td><td>Invitrogen</td><td>NP0321BOX</td><td>SDS-PAGE gel</td></tr><tr><td>4X loading buffer</td><td>Novex</td><td>NP0008</td><td>Protein loading dye concentrate</td></tr><tr><td>50 bp ladder</td><td>New England Biolabs</td><td>N3236</td><td>Reference ladder for excising region of interest from the amplified cDNA library</td></tr><tr><td>50% PEG</td><td>NEB</td><td>B100045</td><td>For the L5 linker ligation</td></tr><tr><td>6% TBE gel</td><td>Invitrogen</td><td>EC6265BOX</td><td>For separation and purification of the cDNA library</td></tr><tr><td>Acetone</td><td>ACROS Organics</td><td>423245000</td><td>Washing of TCA-precipitated proteins</td></tr><tr><td>anti-FLAG beads</td><td>Sigma Aldrich</td><td>M8823-1ML</td><td>For purifcation of FLAG-tagged RBPs</td></tr><tr><td>ATP (100 mM)</td><td>Thermo Fisher Scientific</td><td>R0441</td><td>For ligation of the L5 linker onto the 5&#039; end of captured RNAs</td></tr><tr><td>Beta-mercaptoethanol</td><td>Sigma Aldrich</td><td>M3148-100ML</td><td>Buffer component</td></tr><tr><td>Biomax MS intensifying screen</td><td>Sigma Aldrich</td><td>Z363162-1EA</td><td>For intensifying the autoradiogram signal</td></tr><tr><td>Chloroform</td><td>Thermo Fisher Scientific</td><td>1010219</td><td>For phenol-chloroform extraction following RNA purification</td></tr><tr><td>cOmplete EDTA-free protease inhibitor cocktail</td><td>Roche</td><td>11873580001</td><td>For inhibition of cellular proteases after lysis</td></tr><tr><td>Complete supplement mixture -TRP</td><td>Formedium</td><td>DCS0149</td><td>For preparation of synthetic defined medium</td></tr><tr><td>Costar Spin-X 0.22 &amp;micro;m filters</td><td>Sigma Aldrich</td><td>CLS8160</td><td>For isolating the excised cDNAs following gel extraction</td></tr><tr><td>DNase RQ1</td><td>Promega</td><td>M6101</td><td>For DNA digest following cell lysis</td></tr><tr><td>dNTPs (10 mM)</td><td>Sigma Aldrich</td><td>4638956001</td><td>For reverse transcription and PCR</td></tr><tr><td>Ethanol</td><td>Thermo Fisher Scientific</td><td>10041814</td><td>For phenol-chloroform extraction following RNA purification and DNA precipitation</td></tr><tr><td>Ethylenediaminetetraacetic acid</td><td>Invitrogen</td><td>AM9261</td><td>For protease K buffer</td></tr><tr><td>Exonuclease I</td><td>New England Biolabs</td><td>M0293</td><td>For degradation of primers following PCR</td></tr><tr><td>Glass microfiber filters</td><td>Whatman</td><td>1823-010</td><td>For isolating the excised cDNAs following gel extraction</td></tr><tr><td>Glucose</td><td>Formedium</td><td>GLU03</td><td>For preparation of glucose-containing, synthetic defined medium</td></tr><tr><td>Glycogen (20 mg/mL)</td><td>Roche</td><td>10901393001</td><td>Precipitation of proteins, RNA and DNA</td></tr><tr><td>GST-TEV</td><td>Homemade</td><td></td><td>Construct and purification protocol is available upon request</td></tr><tr><td>Guanidium hydrochloroide</td><td>Thermo Fisher Scientific</td><td>10071503</td><td>Required for pulldown denaturing conditions and washing buffer</td></tr><tr><td>IgG beads</td><td>GE Healthcare</td><td>17-0969-01</td><td>For purification of protein A-tagged RBPs</td></tr><tr><td>Imidazole</td><td>Sigma Aldrich</td><td>I2399-100G</td><td>For elution of captured proteins from Nickel beads</td></tr><tr><td>Isoamyl alcohol</td><td>Thermo Fisher Scientific</td><td>A393-500</td><td>For phenol-chloroform extraction following RNA purification</td></tr><tr><td>Luna Universal One-Step RT-qPCR</td><td>NEB</td><td>E3005S</td><td>For qPCR of the cDNA in order to calculate required number of PCR cycles</td></tr><tr><td>Magnesium chloride</td><td>Fluka Analytical</td><td>63020-1L</td><td>For PNK buffer</td></tr><tr><td>Membrane filters</td><td>Millipore</td><td>AAWP09000 for yeast or HAWP09000 for bacteria</td><td>For vacuum filtration of cells</td></tr><tr><td>Micro bio-spin columns</td><td>Biorad</td><td>732-6204</td><td>For collecting eluate after gel extraction</td></tr><tr><td>Ni-NTA beads</td><td>Qiagen</td><td>30210</td><td>For secondary protein capture</td></tr><tr><td>NP-40</td><td>Sigma Aldrich</td><td>I8896-100ML</td><td>Buffer component</td></tr><tr><td>Pfu polymerase</td><td>Promega</td><td>M7741</td><td>For amplification of the cDNA library</td></tr><tr><td>Phenol</td><td>Sigma Aldrich</td><td>P4682-400ML</td><td>For phenol-chloroform extraction following RNA purification</td></tr><tr><td>Pierce spin columns</td><td>Thermo Fisher Scientific</td><td>69725</td><td>For on-column enzymatic reactions</td></tr><tr><td>Protease K</td><td>Roche</td><td>3115887001</td><td>For degradation of the RBP following gel extraction</td></tr><tr><td>Quartz Petri dish</td><td>UVO3</td><td>N/A</td><td>For cross-linking of adherent cells. Available from https://www.vari-x-link.com for 400 GBP</td></tr><tr><td>Radiography films</td><td>Amersham</td><td>28906843</td><td>For autoradiography visualisation</td></tr><tr><td>RNAClean XP beads</td><td>Beckmann</td><td>A63987</td><td>SPRI beads for clean up of RNAs and cDNAs</td></tr><tr><td>RNase H</td><td>New England Biolabs</td><td>M0297</td><td>For degradation of RNAs following reverse transcription</td></tr><tr><td>RNase-It</td><td>Agilent</td><td>400720</td><td>For RNA digestion</td></tr><tr><td>rRNasin</td><td>Promega</td><td>N2511</td><td>For inhibition of any contaminating RNases during enzymatic reaction</td></tr><tr><td>Sodium acetate</td><td>Sigma Aldrich</td><td>S2889-1KG</td><td>For phenol-chloroform extraction following RNA purification and DNA precipitation</td></tr><tr><td>Sodium chloride</td><td>Thermo Fisher Scientific</td><td>7647-14-5</td><td>Buffer component</td></tr><tr><td>Sodium deoxycholate</td><td>Sigma Aldrich</td><td>D6750-100G</td><td>Buffer component in mammalian cell lysis</td></tr><tr><td>Sodium dodecylsulfate</td><td>Sigma Aldrich</td><td>L3771-1KG</td><td>For protease K buffer</td></tr><tr><td>SUPERase-In</td><td>Invitrogen</td><td>AM2694</td><td>For inhibition of cellular RNases after lysis</td></tr><tr><td>SuperScript IV</td><td>Thermo Fisher Scientific</td><td>18090010</td><td>For reverse transcription</td></tr><tr><td>T4 PNK</td><td>New England Biolabs</td><td>M0201</td><td>For radiolabelling the 5&#039; end of the RNA</td></tr><tr><td>T4 RNA ligase 1</td><td>New England Biolabs</td><td>M0204</td><td>For ligation of the L5 adaptor onto the RNA 5&#039; end</td></tr><tr><td>T4 RNase ligase 2, truncated K222Q</td><td>NEB</td><td>M0351S</td><td>For ligation of the App_PE linker onto the 3&#039; end of captured RNAs</td></tr><tr><td>TBE buffer (10X)</td><td>Invitrogen</td><td>15581-028</td><td>For running TBE gels</td></tr><tr><td>TEV protease</td><td>Homemade</td><td></td><td>For eluting captured proteins following FLAG capture</td></tr><tr><td>Thermosensitive alkaline phosphatase</td><td>Promega</td><td>M9910</td><td>For 5&#039; and 3&#039; dephosphorylation of RNAs</td></tr><tr><td>Trichloroacetic acid (100%)</td><td>Sigma Aldrich</td><td>T0699-100ML</td><td>For precipitation of RBP-RNA complexes</td></tr><tr><td>Tris hydrochloride</td><td>Invitrogen</td><td>15504-020</td><td>Buffer component</td></tr><tr><td>Triton X-100</td><td>Sigma Aldrich</td><td>T8787-100ML</td><td>Buffer component in mammalian cell lysis</td></tr><tr><td>Vari Filter</td><td>UVO3</td><td>N/A</td><td>Device for vacuum harvesting cells. Available from https://www.vari-x-link.com for 100 GBP</td></tr><tr><td>Vari-X-Linker</td><td>UVO3</td><td>N/A</td><td>Cross-linker for cross-linking cells. Available from https://www.vari-x-link.com for 16,000 GBP</td></tr><tr><td>Yeast nitrogen base</td><td>Formedium</td><td>CYN0410</td><td>For preparation of synthetic defined medium</td></tr><tr><td>Zirconia beads</td><td>Thistle</td><td>11079105Z for yeast or 11079101Z for bacteria</td><td>For cell lysis via bead beating</td></tr></tbody></table>

monitoring protein-rna interaction dynamics in vivo at high temporal resolution using &#967;crac

The interaction between RNA-binding proteins (RBPs) and their RNA substrates exhibits fluidity and complexity. Within its lifespan, a single RNA can be bound by many different RBPs that will regulate its production, stability, activity, and degradation. As such, much has been done to understand the dynamics that exist between these two types of molecules. A particularly important breakthrough came with the emergence of &#8216;cross-linking and immunoprecipitation&#8217; (CLIP). This technique allowed stringent investigation into which RNAs are bound by a particular RBP. In short, the protein of interest is UV cross-linked to its RNA substrates in vivo, purified under highly stringent conditions, and then the RNAs covalently cross-linked to the protein are converted into cDNA libraries and sequenced. Since its conception, many derivative techniques have been developed in order to make CLIP amenable to particular fields of study. However, cross-linking using ultraviolet light is notoriously inefficient. This results in extended exposure times that make the temporal study of RBP-RNA interactions impossible. To overcome this issue, we recently designed and built much-improved UV irradiation and cell harvesting devices. Using these new tools, we developed a protocol for time-resolved analyses of RBP-RNA interactions in living cells at high temporal resolution: Kinetic CRoss-linking and Analysis of cDNAs (&#967;CRAC). We recently used this technique to study the role of yeast RBPs in nutrient stress adaptation. This manuscript provides a detailed overview of the &#967;CRAC method and presents recent results obtained with the Nrd1 RBP.

To demonstrate the efficacy of the &#967;CRAC method, a time-course experiment with yeast strains expressing an HTP-tagged Nrd1 protein was performed. A detailed schematic representation describing how the method works is provided in Figure 1. Like Nab3, Nrd1 is involved in nuclear RNA decay of a variety of RNA transcripts37. Previous work from the Corden lab suggested that Nrd1 binding to its RNA targets changes significantly when cells are subjected to glucose starvation28,38. As such, cells growing exponentially in medium containing glucose (SD-TRP) were shifted to the same medium without glucose (S-TRP) over a time-course to monitor dynamic changes in Nrd1-RNA interactions. Samples were taken and cross-linked in the Vari-X-linker chamber (Figure 3A) before the shift and then after 1, 2, 4, 8, 14, and 20 min. The medium used for cell growth was deliberately deficient in tryptophan to reduce UV absorption by this aromatic amino acid. Note that it is best to use synthetic medium that is filter sterilized as&#160;because autoclaving the medium can lead to caramelization of the sugars. This then&#160;reduces the cross-linking efficiency.
Figure 4A shows a representative autoradiograph from a &#967;CRAC experiment. Note that in this example, the samples were not pooled together. Instead, each was run individually on the gel. This is recommended for initial experimental tests to show that the protein cross-links effectively to RNA at all of the tested timepoints. A particularly intense signal was observed at the expected molecular weight of the RBP, representing the protein bound to very short, radiolabeled RNAs not amenable for sequencing. Therefore, the smeary signal above this band, which is the protein crosslinked to longer RNA fragments, was isolated. The fragment was cut from just above the protein band plus around 30 kDa. Figure 4B shows an autoradiogram following excision, with the protein cross-linked to short RNAs left in the gel and the previously smeary signal now excised.
After reverse transcription, the cDNA library must be amplified using PCR. However, overamplification of the library must be avoided as this can introduce bias towards sequences preferentially amplified by the polymerase and generate PCR artefacts. Overamplified libraries also contain a large number of duplicate sequences that waste reads on the sequencer. In order to calculate the ideal number of PCR cycles for amplification of the final library, an aliquot of the cDNA was amplified through qPCR using the P5 and BC oligonucleotides. The first cycle at which the library reached peak fluorescence was chosen as the PCR cycle count. Figure 4C gives an example of a qPCR from a typical cDNA library, which yielded a peak cycle count of 16. This value was then used for the final &#967;CRAC PCR. In order to process the sequenced data, we used software previously developed in our lab (pyCRAC) and the corresponding pipeline for analysis of the kinetic CRAC data (Nues et al., 2017; https://git.ecdf.ed.ac.uk/sgrannem/pycrac, https://bitbucket.org/sgrann/kinetic_crac_pipeline/src/default/). These open source software tools enable demultiplexing and trimming of the data, removal of PCR duplicates, identification of statistically significant peaks, cluster reads into contiguous sequences, and identify binding motifs39. Further details of how these tools operate are found on their respective webpages.
We also started to develop a &#967;CRAC protocol for mammalian cells. The majority of mammalian cell lines are grown as a monolayer and the tray in our crosslinker with the UV-permeable bag is not suitable for experiments with adherent cells. To overcome this problem, we developed a stage where users can UV irradiate 1&#8211;2 Petri dishes (150 mm diameter and 25 mm in depth) with adherent cells (Figure 3B). As a first test, the efficiency of the cross-linker for mammalian cells was measured through cross-linking and capture of stably tagged GFP-RBM7 using anti-GFP antibodies and a traditional CLIP-based purification. As shown in Figure 5A, the cross-linker was able to recover protein-RNA complexes from mammalian cells grown as a monolayer using 254 nm UV irradiation at efficiencies comparable to a widely used UV irradiation device. However, standard cell culture plasticware normally used for UV cross-linking experiments is impenetrable to 254 nm UV. Therefore, in our cross-linker the cells would only receive irradiation from the upper bank of UV lamps. To overcome this, we developed a UV-permeable quartz Petri dish for cell growth and cross-linking. Use of the quartz cultureware displayed robust recovery of protein-RNA complexes with as few as 2 s of UV irradiation (Figure 5B). When combined with RBP capture methods for mammalian cells such as CLIP technologies, these short cross-linking times are amenable with timecourses to recover spatiotemporal RNA-binding profiles of RBPs in response to genotoxic stresses or rapid depletions of protein factors, or in parallel with transcriptional or cell cycle synchronization.
Figure 6 shows several examples of the Nrd1 data processed by the &#967;CRAC pipeline. This figure was prepared using the bedgraph files generated by the pipeline and the python GenomeBrowser package (https://pypi.org/project/GenomeBrowser/1.6.3/), which we designed to simplify making publication-quality genome browser images of the data. The grey rectangles represent genomic regions that expressed noncoding RNAs, such as the cryptic unstable transcript (CUTs), stable uncharacterized transcripts (SUTs)40, and Xrn1-sensitive unstable transcripts (XUTs)41. The data in Figure 6 show that Nrd1 binds to many of these noncoding RNA transcripts, consistent with the idea that this protein is involved in degradation of this class of transcripts42. Figure 6A shows a ~15 kb region on chromosome IV. Here there was a significant increase in binding of Nrd1 to transcripts encoding the high-affinity glucose transporters HXT6 and HXT7, both of which are upregulated during glucose starvation. It is likely that transcription termination by the NNS complex may influence the induction kinetics of these genes during glucose starvation. Figure 6B shows an example of Nrd1 cross-linking to the Imd3 transcript, which is known to be regulated by Nab343. In this case the data demonstrated a significant reduction in binding upon glucose starvation. Previous work showed&#160;decreased binding of Nab3 to the Tye7 transcript during glucose starvation44. Consistent with this observation,the &#967;CRAC data suggest that binding of Nrd1 decreased during glucose starvation and Nrd1 cross-linking to Tye7 was at its lowest after 8 min of stress (Figure 4C). However, it appears that this effect was only transient, because after 14 min of glucose starvation, Nrd1 binding went back to starting levels.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61027/61027fig1v2.jpg" /> 
Figure 1: Schematic representation of the &#967;CRAC protocol. Tagged strains were grown until the desired density. RBP indicates RNA-binding protein. Afterwards, a reference sample was taken and cross-linked with 254 nm UV light. The remaining cells were harvested by filtration and then rapidly shifted to the stress-inducing medium. For the &#967;CRAC experiment described here, samples were taken and cross-linked 1, 2, 4, 8, 14, and 20 min after the shift (1). The RBP of interest was then purified using a highly stringent two-step affinity purification (2). Next, the captured cross-linked RNAs were partially digested with RNases, radiolabeled at the 5&#8217; end and adapters were ligated onto them (3). The 5&#8217; adapters contained unique &#8220;in-read&#8221; barcode sequences so that the individual samples could be separated bioinformatically after sequencing. The RBP-RNA complexes were then eluted, pooled, and precipitated together (4), resolved by SDS-PAGE and visualized through autoradiography (5). Subsequently, a single gel slice containing the radioactive signal just above the main band, illustrated with dashed red box in the autoradiography image, was cut from the gel (5). The gel slices were treated with protease K and the RNA was subsequently extracted (6), converted into cDNAs, and amplified through PCR (7). The PCR step introduced additional barcodes (yellow block introduced by P7 oligo) so that many libraries could be multiplexed into a single lane. <a href="https://www.jove.com/files/ftp_upload/61027/61027fig1v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61027/61027fig2v2.jpg" /> 
Figure 2: Cross-linking and vacuum filtration. (A) The cross-linker. The cell suspension is poured into a funnel located in the top right of the machine (also see Figure 3A for a close-up) and held in a UV-transparent bag located in the middle tray. This bag is flanked by two shutters that remain closed until the user instructs the machine to start the irradiation step. The cells are irradiated with UV light from the trays both above and below. The machine comes supplied with 254 and 365 nm UV lamps, with the latter being applicable for PAR-CLIP experiments. The machine is operated through a touchscreen panel located on the top right which allows one to control UV dosage or exposure time. (B) Following cross-linking, the cells are drained from the left-hand side of the machine. Cell suspensions are recovered through vacuum and drained into a glass flask where they can be subsequently poured into a vacuum filtration device for harvesting. (C) Vacuum filtration devices. These are opened and closed via a clip and a filter is inserted between it. Four filtration devices were used in parallel for very short time series to not lose any time as a result of changing filters. (D) Following filtration, the media supernatant was drained into flasks for subsequent disposal. Valves were installed below the vacuum filtration devices to maintain the vacuum in the system when the filter is removed. <a href="https://www.jove.com/files/ftp_upload/61027/61027fig2v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61027/61027fig3v2.jpg" /> 
Figure 3: Cross-linking suspended vs. adherent cells. (A) The cross-linker with the Vari-X-linker chamber for suspension cells. The cell culture is poured into the sample inlet (funnel) located in the top right of the tray. (B) Tray that can hold plastic or quartz Petri dishes for cross-linking adherent cells or small volumes of suspension cells. <a href="https://www.jove.com/files/ftp_upload/61027/61027fig3v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61027/61027fig4v2.jpg" /> 
Figure 4: Library preparation. (A) Example of an autoradiogram from a Nrd1-HTP &#967;CRAC experiment. The strong, concentrated signal represents the protein crosslinked to very short RNAs, while the smear above represents the protein cross-linked to RNAs of sufficient length for sequencing. (B) The smear was excised as shown in an autoradiogram taken after gel excision. (C) A representative qPCR from a &#967;CRAC cDNA library. In this example, maximum amplification of the cDNA was reached at 16 cycles. Thus, 16 cycles were used for the final amplification. The error bar represents the standard deviation of three technical qPCR replicates. (D) Example of a phosphorimage from a cDNA library on a 6% TBE gel. (E) cDNA length and quality analysis from a chip-based capillary electrophoresis. <a href="https://www.jove.com/files/ftp_upload/61027/61027fig4v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/61027/61027fig5v2.jpg" /> 
Figure 5: High RNase test iCLIP experiment to test crosslinking in mammalian cells. Shown are autoradiograms from GFP-RBM7 iCLIP experiments which tested the efficiency of RNP recovery across various cross-linking energies. Immunoprecipitations were performed using anti-GFP antibodies coupled to magnetic beads on cross-linked cells which stably expressed GFP-RBM7. Immunoprecipitates were incubated with high concentrations of RNase I in order to trim associated RNAs to short, uniform lengths. RNPs were visualized by 32P labelling and SDS-PAGE and migrate as a defined band, close to the migration of the non-cross-linked protein. Quantification indicates the results of densitometric analyses of radiolabeled RBM7-RNA signal normalized to the anti-GFP western blot signal. (A) Cross-linking time-course of the commonly used UVP cross-linker versus our cross-linker (Vari-X-linker; VxL). (B) Cross-linking time-course of our cross-linker on quartz (left) and plastic (right) cultureware. <a href="https://www.jove.com/files/ftp_upload/61027/61027fig5v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/61027/61027fig6v2.jpg" /> 
Figure 6: Example genome browser plots showing the power of &#967;CRAC to show differential, temporal binding of Nrd1 to its targets. Each box shows plots for individual genomic regions. The arrows indicate on which strand the genes are encoded (left pointing arrow = minus strand; right pointing arrow = plus strand). The timepoints (min) are indicated by t0, t1, t2, etc on the y-axes of each subplot. Roman numerals indicating the chromosomes and the coordinates are shown. (A) Upon glucose deprivation, Nrd1 binds two high-affinity glucose transporters, HXT6 and HXT7, which are both upregulated in this condition. (B) Nrd1 is observed to bind to Imd3, an already validated target of Nab344, with reducing intensity following glucose starvation. (C) Nrd1 binding of Tye7 exhibits a dynamic and transient nature, decreasing after glucose starvation to a minimum after 8 min of stress. However, binding subsequently returns to basal levels after 14 min. Reads were normalized to &#8220;reads per million&#8221; (RPM; y-axis). Grey boxes indicate regions encoding noncoding RNAs. <a href="https://www.jove.com/files/ftp_upload/61027/61027fig6v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Monitoring Protein-RNA Interaction Dynamics In Vivo at High Temporal Resolution Using Ï‡CRAC at JoVE.com

Monitoring Protein-RNA Interaction Dynamics In Vivo at High Temporal Resolution Using &#967;CRAC

Kinetic cross-linking and analysis of cDNA is a method that allows investigation of the dynamics of protein-RNA interactions in living cells at high temporal resolution. Here the protocol is described in detail, including the growth of yeast cells, UV cross-linking, harvesting, protein purification, and next generation sequencing library preparation steps.

The interaction between RNA-binding proteins (RBPs) and their RNA substrates exhibits fluidity and complexity. Within its lifespan, a single RNA can be ...

monitoring-protein-rna-interaction-dynamics-vivo-at-high-temporal

Research

JoVE Journal

Biochemistry

4.9K Views.  University of Edinburgh. Kinetic cross-linking and analysis of cDNA is a method that allows investigation of the dynamics of protein-RNA interactions in living cells at high temporal resolution. Here the protocol is described in detail, including the growth of yeast cells, UV cross-linking, harvesting, protein purification, and next generation sequencing library preparation steps.

Video: Monitoring Protein-RNA Interaction Dynamics In Vivo at High Temporal Resolution Using χCRAC

Novel RNA-Binding Proteins Isolation by the RaPID Methodology

RNA-binding proteins (RBPs) play important roles in every aspect of RNA metabolism and regulation. Their identification is a major challenge in modern biology. Only a few in vitro and in vivo methods enable the identification of RBPs associated with a particular target mRNA. However, their main limitations are the identification of RBPs in a non-cellular environment (in vitro) or the low efficiency isolation of RNA of interest (in vivo). An RNA-binding protein purification and identification (RaPID) methodology was designed to overcome these limitations in yeast and enable efficient isolation of proteins that are associated in vivo. To achieve this, the RNA of interest is tagged with MS2 loops, and co-expressed with a fusion protein of an MS2-binding protein and a streptavidin-binding protein (SBP). Cells are then subjected to crosslinking and lysed, and complexes are isolated through streptavidin beads. The proteins that co-purify with the tagged RNA can then be determined by mass spectrometry. We recently used this protocol to identify novel proteins associated with the ER-associated PMP1 mRNA. Here, we provide a detailed protocol of RaPID, and discuss some of its limitations and advantages.

RNA-protein interactions lie at the heart of many cellular processes. Here, we describe an in vivo method to isolate specific RNA and identify novel proteins that are associated with it. This could shed new light on how RNAs are regulated in the cell.

RNA-binding proteins (RBPs) play important roles in every aspect of RNA metabolism and regulation. Their identification is a major challenge in modern ...

Genetics

Real-time Analysis of Transcription Factor Binding, Transcription, Translation, and Turnover to Display Global Events During Cellular Activation

Upon activation, cells rapidly change their functional programs and, thereby, their gene expression profile. Massive changes in gene expression occur, for example, during cellular differentiation, morphogenesis, and functional stimulation (such as activation of immune cells), or after exposure to drugs and other factors from the local environment. Depending on the stimulus and cell type, these changes occur rapidly and at any possible level of gene regulation. Displaying all molecular processes of a responding cell to a certain type of stimulus/drug is one of the hardest tasks in molecular biology. Here, we describe a protocol that enables the simultaneous analysis of multiple layers of gene regulation. We compare, in particular, transcription factor binding (Chromatin-immunoprecipitation-sequencing (ChIP-seq)), de novo transcription (4-thiouridine-sequencing (4sU-seq)), mRNA processing, and turnover as well as translation (ribosome profiling). By combining these methods, it is possible to display a detailed and genome-wide course of action.
Sequencing newly transcribed RNA is especially recommended when analyzing rapidly adapting or changing systems, since this depicts the transcriptional activity of all genes during the time of 4sU exposure (irrespective of whether they are up- or downregulated). The combinatorial use of total RNA-seq and ribosome profiling additionally allows the calculation of RNA turnover and translation rates. Bioinformatic analysis of high-throughput sequencing results allows for many means for analysis and interpretation of the data. The generated data also enables tracking co-transcriptional and alternative splicing, just to mention a few possible outcomes.
The combined approach described here can be applied for different model organisms or cell types, including primary cells. Furthermore, we provide detailed protocols for each method used, including quality controls, and discuss potential problems and pitfalls.

This protocol describes the combinatorial use of ChIP-seq, 4sU-seq, total RNA-seq, and ribosome profiling for cell lines and primary cells. It enables tracking changes in transcription-factor binding, de novo transcription, RNA processing, turnover and translation over time, and displaying the overall course of events in activated and/or rapidly changing cells.

Upon activation, cells rapidly change their functional programs and, thereby, their gene expression profile. Massive changes in gene expression occur, for ...

mRNA Interactome Capture from Plant Protoplasts

RNA-binding proteins (RBPs) determine the fates of RNAs. They participate in all RNA biogenesis pathways and especially contribute to post-transcriptional gene regulation (PTGR) of messenger RNAs (mRNAs). In the past few years, a number of mRNA-bound proteomes from yeast and mammalian cell lines have been successfully isolated through the use of a novel method called &#34;mRNA interactome capture,&#34; which allows for the identification of mRNA-binding proteins (mRBPs) directly from a physiological environment. The method is composed of in vivo ultraviolet (UV) crosslinking, pull-down and purification of messenger ribonucleoprotein complexes (mRNPs) by oligo(dT) beads, and the subsequent identification of the crosslinked proteins by mass spectrometry (MS). Very recently, by applying the same method, several plant mRNA-bound proteomes have been reported simultaneously from different Arabidopsis tissue sources: etiolated seedlings, leaf tissue, leaf mesophyll protoplasts, and cultured root cells. Here, we present the optimized mRNA interactome capture method for Arabidopsis thaliana leaf mesophyll protoplasts, a cell type that serves as a versatile tool for experiments that include various cellular assays. The conditions for optimal protein yield include the amount of starting tissue and the duration of UV irradiation. In the mRNA-bound proteome obtained from a medium-scale experiment (107 cells), RBPs noted to have RNA-binding capacity were found to be overrepresented, and many novel RBPs were identified. The experiment can be scaled up (109 cells), and the optimized method can be applied to other plant cell types and species to broadly isolate, catalog, and compare mRNA-bound proteomes in plants.

Here, we present an interactome capture protocol applied to Arabidopsis thaliana leaf mesophyll protoplasts. This method critically relies on in vivo UV crosslinking and allows for the isolation and identification of plant mRNA-binding proteins from a physiological environment.

RNA-binding proteins (RBPs) determine the fates of RNAs. They participate in all RNA biogenesis pathways and especially contribute to post-transcriptional ...

An Assay for Quantifying Protein-RNA Binding in Bacteria

In the initiation step of protein translation, the ribosome binds to the initiation region of the mRNA. Translation initiation can be blocked by binding of an RNA binding protein (RBP) to the initiation region of the mRNA, which interferes with ribosome binding. In the presented method, we utilize this blocking phenomenon to quantify the binding affinity of RBPs to their cognate and non-cognate binding sites. To do this, we insert a test binding site in the initiation region of a reporter mRNA and induce the expression of the test RBP. In the case of RBP-RNA binding, we observed a sigmoidal repression of the reporter expression as a function of RBP concentration. In the case of no-affinity or very low affinity between binding site and RBP, no significant repression was observed. The method is carried out in live bacterial cells, and does not require expensive or sophisticated machinery. It is useful for quantifying and comparing between the binding affinities of different RBPs that are functional in bacteria to a set of designed binding sites. This method may be inappropriate for binding sites with high structural complexity. This is due to the possibility of repression of ribosomal initiation by complex mRNA structure in the absence of RBP, which would result in lower basal reporter gene expression, and thus less-observable reporter repression upon RBP binding.

In this method, we quantify the binding affinity of RNA binding proteins (RBPs) to cognate and non-cognate binding sites using a simple, live, reporter assay in bacterial cells. The assay is based on repression of a reporter gene.

In the initiation step of protein translation, the ribosome binds to the initiation region of the mRNA. Translation initiation can be blocked by binding ...

Capture and Identification of RNA-binding Proteins by Using Click Chemistry-assisted RNA-interactome Capture (CARIC) Strategy

A comprehensive identification of RNA-binding proteins (RBPs) is key to understanding the posttranscriptional regulatory network in cells. A widely used strategy for RBP capture exploits the polyadenylation [poly(A)] of target RNAs, which mostly occurs on eukaryotic mature mRNAs, leaving most binding proteins of non-poly(A) RNAs unidentified. Here we describe the detailed procedures of a recently reported method termed click chemistry-assisted RNA-interactome capture (CARIC), which enables the transcriptome-wide capture of both poly(A) and non-poly(A) RBPs by combining the metabolic labeling of RNAs, in vivo UV cross-linking, and bioorthogonal tagging.

Capture and Identification of RNA-binding Proteins by Using Click Chemistry-assisted RNA-interactome Capture CARIC Strategy

A detailed protocol for applying the click chemistry-assisted RNA-interactome capture (CARIC) strategy to identify proteins binding to both coding and noncoding RNAs is presented.

A comprehensive identification of RNA-binding proteins (RBPs) is key to understanding the posttranscriptional regulatory network in cells. A widely used ...

Studying RNA Interactors of Protein Kinase RNA-Activated during the Mammalian Cell Cycle

Protein kinase RNA-activated (PKR) is a member of the innate immune response proteins and recognizes the double-stranded secondary structure of viral RNAs. When bound to viral double-stranded RNAs (dsRNAs), PKR undergoes dimerization and subsequent autophosphorylation. Phosphorylated PKR (pPKR) becomes active and induces phosphorylation of the alpha subunit of eukaryotic initiation factor 2 (eIF2&#945;) to suppress global translation. Increasing evidence suggests that PKR can be activated under physiological conditions such as during the cell cycle or under various stress conditions without infection. However, our understanding of the RNA activators of PKR is limited due to the lack of a standardized experimental method to capture and analyze PKR-interacting dsRNAs. Here, we present an experimental protocol to specifically enrich and analyze PKR bound RNAs during the cell cycle using HeLa cells. We utilize the efficient crosslinking activity of formaldehyde to fix PKR-RNA complexes and isolate them via immunoprecipitation. PKR co-immunoprecipitated RNAs can then be further processed to generate a high-throughput sequencing library. One major class of PKR-interacting cellular dsRNAs is mitochondrial RNAs (mtRNAs), which can exist as intermolecular dsRNAs through complementary interaction between the heavy-strand and the light-strand RNAs. To study the strandedness of these duplex mtRNAs, we also present a protocol for strand-specific qRT-PCR. Our protocol is optimized for the analysis of PKR-bound RNAs, but it can be easily modified to study cellular dsRNAs or RNA-interactors of other dsRNA binding proteins.

We present experimental approaches for studying RNA-interactors of double-stranded RNA binding protein kinase RNA-activated (PKR) during the mammalian cell cycle using HeLa cells. This method utilizes formaldehyde to crosslink RNA-PKR complexes and immunoprecipitation to enrich PKR-bound RNAs. These RNAs can be further analyzed through high-throughput sequencing or qRT-PCR.

Protein kinase RNA-activated (PKR) is a member of the innate immune response proteins and recognizes the double-stranded secondary structure of viral ...

A Rapid High-throughput Method for Mapping Ribonucleoproteins (RNPs) on Human pre-mRNA

Sequencing RNAs that co-immunoprecipitate (co-IP) with RNA binding proteins has increased our understanding of splicing by demonstrating that binding location often influences function of a splicing factor.  However, as with any sampling strategy the chance of identifying an RNA bound to a splicing factor is proportional to its cellular abundance.  We have developed a novel in vitro approach for surveying binding specificity on otherwise transient pre-mRNA. This approach utilizes a specifically designed oligonucleotide pool that tiles across introns, exons, splice junctions, or other pre-mRNA.  The pool is subjected to some kind of molecular selection. Here, we demonstrate the method by separating the oligonucleotide into a bound and unbound fraction and utilize a two color array strategy to record the enrichment of each oligonucleotide in the bound fraction. The array data generates high-resolution maps with the ability to identify sequence-specific and structural determinates of ribonucleoprotein (RNP) binding on pre-mRNA.  A unique advantage to this method is its ability to avoid the sampling bias towards mRNA associated with current IP and SELEX techniques, as the pool is specifically designed and synthesized from pre-mRNA sequence. The flexibility of the oligonucleotide pool is another advantage since the experimenter chooses which regions to study and tile across, tailoring the pool to their individual needs.  Using this technique, one can assay the effects of polymorphisms or mutations on binding on a large scale or clone the library into a functional splicing reporter and identify oligonucleotides that are enriched in the included fraction.  This novel in vitro high-resolution mapping scheme provides a unique way to study RNP interactions with transient pre-mRNA species, whose low abundance makes them difficult to study with current in vivo techniques.  



A Rapid High-throughput Method for Mapping Ribonucleoproteins RNPs on Human pre-mRNA

Due to the transient nature of pre-mRNA, it can be difficult to isolate and study in vivo. Here, we present a novel in vitro approach to investigate RNA-protein interactions using a synthetic oligo pool that tiles across selected regions of pre-mRNA.

Sequencing RNAs that co-immunoprecipitate (co-IP) with RNA binding proteins has increased our understanding of splicing by demonstrating that binding ...

Biology

Monitoring Protein-Ligand Interactions in Human Cells by Real-Time Quantitative In-Cell NMR using a High Cell Density Bioreactor

In-cell NMR is a unique approach to observe the structural and dynamic properties of biological macromolecules at atomic resolution directly in living cells. Protein folding, chemical modifications, and conformational changes induced by ligand binding can be observed. Therefore, this method has great potential in the context of drug development. However, the short lifetime of human cells confined in the NMR spectrometer limits the application range of in-cell NMR. To overcome this issue, NMR bioreactors are employed that can greatly improve the cell sample stability over time and, importantly, enable the real-time recording of in-cell NMR spectra. In this way, the evolution of processes such as ligand penetration and binding to the intracellular protein target can be monitored in real time. Bioreactors are often limited by low cell viability at high cell numbers, which results in a trade-off between the overall sensitivity of the experiment and cell viability. We recently reported an NMR bioreactor that maintains a high number of human cells metabolically active for extended periods of time, up to 72 h. This setup was applied to monitor protein-ligand interactions and protein chemical modification. We also introduced a workflow for quantitative analysis of the real-time NMR data, based on multivariate curve resolution. The method provides concentration profiles of the chemical species present in the cells as a function of time, which can be further analyzed to obtain relevant kinetic parameters. Here we provide a detailed description of the NMR bioreactor setup and its application to monitoring protein-ligand interactions in human cells.

This protocol describes the setup of an NMR bioreactor to keep encapsulated human cells viable for up to 72 h, followed by time-resolved in-cell NMR data acquisition and analysis. The methodology is applied to monitor intracellular protein-ligand interactions in real time.

In-cell NMR is a unique approach to observe the structural and dynamic properties of biological macromolecules at atomic resolution directly in living ...

Chemistry

iCLIP - Transcriptome-wide Mapping of Protein-RNA Interactions with Individual Nucleotide Resolution

The unique composition and spatial arrangement of RNA-binding proteins (RBPs) on a transcript guide the diverse aspects of post-transcriptional regulation1. Therefore, an essential step towards understanding transcript regulation at the molecular level is to gain positional information on the binding sites of RBPs2.
Protein-RNA interactions can be studied using biochemical methods, but these approaches do not address RNA binding in its native cellular context. Initial attempts to study protein-RNA complexes in their cellular environment employed affinity purification or immunoprecipitation combined with differential display or microarray analysis (RIP-CHIP)3-5. These approaches were prone to identifying indirect or non-physiological interactions6. In order to increase the specificity and positional resolution, a strategy referred to as CLIP (UV cross-linking and immunoprecipitation) was introduced7,8. CLIP combines UV cross-linking of proteins and RNA molecules with rigorous purification schemes including denaturing polyacrylamide gel electrophoresis. In combination with high-throughput sequencing technologies, CLIP has proven as a powerful tool to study protein-RNA interactions on a genome-wide scale (referred to as HITS-CLIP or CLIP-seq)9,10. Recently, PAR-CLIP was introduced that uses photoreactive ribonucleoside analogs for cross-linking11,12.
Despite the high specificity of the obtained data, CLIP experiments often generate cDNA libraries of limited sequence complexity. This is partly due to the restricted amount of co-purified RNA and the two inefficient RNA ligation reactions required for library preparation. In addition, primer extension assays indicated that many cDNAs truncate prematurely at the crosslinked nucleotide13. Such truncated cDNAs are lost during the standard CLIP library preparation protocol. We recently developed iCLIP (individual-nucleotide resolution CLIP), which captures the truncated cDNAs by replacing one of the inefficient intermolecular RNA ligation steps with a more efficient intramolecular cDNA circularization (Figure 1)14. Importantly, sequencing the truncated cDNAs provides insights into the position of the cross-link site at nucleotide resolution. We successfully applied iCLIP to study hnRNP C particle organization on a genome-wide scale and assess its role in splicing regulation14.

The spatial arrangement of RNA-binding proteins on a transcript is a key determinant of post-transcriptional regulation. Therefore, we developed individual-nucleotide resolution UV crosslinking and immunoprecipitation (iCLIP) that allows precise genome-wide mapping of the binding sites of an RNA-binding protein.

The unique composition and spatial arrangement of RNA-binding proteins (RBPs) on a transcript guide the diverse aspects of post-transcriptional ...

Single-molecule Imaging of Gene Regulation In vivo Using Cotranslational Activation by Cleavage (CoTrAC)

We describe a fluorescence microscopy method, Co-Translational Activation by Cleavage (CoTrAC) to image the production of protein molecules in live cells with single-molecule precision without perturbing the protein's functionality. This method makes it possible to count the numbers of protein molecules produced in one cell during sequential, five-minute time windows. It requires a fluorescence microscope with laser excitation power density of ~0.5 to 1 kW/cm2, which is sufficiently sensitive to detect single fluorescent protein molecules in live cells. The fluorescent reporter used in this method consists of three parts: a membrane targeting sequence, a fast-maturing, yellow fluorescent protein and a protease recognition sequence. The reporter is translationally fused to the N-terminus of a protein of interest. Cells are grown on a temperature-controlled microscope stage. Every five minutes, fluorescent molecules within cells are imaged (and later counted by analyzing fluorescence images) and subsequently photobleached so that only newly translated proteins are counted in the next measurement.
Fluorescence images resulting from this method can be analyzed by detecting fluorescent spots in each image, assigning them to individual cells and then assigning cells to cell lineages. The number of proteins produced within a time window in a given cell is calculated by dividing the integrated fluorescence intensity of spots by the average intensity of single fluorescent molecules. We used this method to measure expression levels in the range of 0-45 molecules in single 5 min time windows. This method enabled us to measure noise in the expression of the &lambda; repressor CI, and has many other potential applications in systems biology.

Single-molecule Imaging of Gene Regulation In vivo Using Cotranslational Activation by Cleavage CoTrAC

We describe a fluorescence microscopy method, Co-Translational Activation by Cleavage (CoTrAC), to image the production of protein molecules in live cells with single-molecule precision without perturbing the protein's functionality. This method has been used to follow the stochastic expression dynamics of a transcription factor, the &lambda; repressor CI 1.

We describe a fluorescence microscopy method, Co-Translational Activation by Cleavage (CoTrAC) to image the production of protein molecules in live cells ...

Monitoring Protein-RNA Interaction Dynamics In Vivo at High Temporal Resolution Using χCRAC

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