This work was supported by the National Institutes of Health [R01GM121847]; the Memorial Sloan Kettering Cancer Center (MSKCC) Support Grant/Core Grant (P30 CA008748); and MSKCC Functional Genomics Initiative.

The authors have nothing to disclose.

In comparison to typical in vitro TIRF single-molecule experiments, single-molecule imaging with the assay described here is additionally complex due to the use of cell extract and a high concentration of fluorescently labeled antibody. Compared to the more common practice of one round of surface PEGylation, a second round of PEGylation (step 2) greatly reduces nonspecific antibody binding to detection surface15. The high concentration of diffusing fluorescent antibodies causes an extremely high fluorescent background that masks single-molecule detection of antibody binding. To reduce this fluorescent background, the laser incident angle is increased well above the critical angle (step 3.4.). Antibody detection is also diminished by excessive fluorophore instability, which can limit the capability to quantify dwell time. When encountering this problem, replace the fluorophore with a more photostable fluorophore, such as the ones conjugated to a triplet-state quencher47, or lower the laser illumination intensity. Although oxygen scavenging systems are commonly used in in vitro single-molecule experiments to suppress fluorophore photobleaching48,49, the assay described here can provide a very generous detection window without implementing any oxygen scavenging systems15. Furthermore, oxygen scavenging systems can greatly reduce eukaryotic cell-free translation efficiency. Therefore, oxygen scavenging systems should be carefully evaluated prior to their use with this assay.
It is important to point out that small variations in reagent preparation are detected by the assay due to its single-molecule resolution. For example, replicate translation extract preparations can display different activities that may or may not be detectable by bulk methods. Furthermore, freeze-thaw cycles of translation extract and other reagents can rapidly impair translation activity. We recommend doing small-scale pilot experiments with readily accessible material to test conditions before planning for a systematic study. For a systematic study, large-scale preparations of translation reagents, single-use aliquots for freeze/thaw-sensitive reagents, and consistency in performing protocol steps are recommended. Translation extract that is flash frozen in liquid nitrogen and stored at -80 &#730;C shows minimal activity loss for at least two years. Application of these measures can effectively suppress experimental variations and increase the robustness of the single-molecule assay.
As this method combines existing bulk cell-free translation systems and in vitro single-molecule imaging techniques, troubleshooting of generic problems in these two areas are not discussed here. Examples of such generic issues include low quality preparations of reporter mRNA or PEGylated coverslip and low translation activity of cell extract. Here, we will focus on issues specific for this method. In addition to the several technical issues that have been discussed in the protocol, another potential problem may be low or no antibody binding in the flow channel for a translation condition that is expected to have good activity. There are several possibilities to troubleshoot. The efficiency of biotinylated mRNA immobilization to the detection surface can be assessed by annealing complementary and fluorophore-conjugated DNA oligomers to the immobilized mRNA and imaging the fluorescent DNA:RNA duplexes. The quality of the translation mix and biotinylated reporter mRNAs can be checked by running the bulk translation assay. Furthermore, after carrying out a translation reaction in an mRNA-immobilized flow channel, the translation reaction solution can be pipetted out of the channel and used in a reporter activity assay to confirm the occurrence of in-channel translation. If necessary, an excessively high mRNA surface density can be used for the in-channel translation reaction to allow easier detection by the reporter activity assay.
The major limitation of this assay is its resolution for initiation kinetics. In this assay, the direct experimental output, first arrival time, contains both the first-round initiation time and the peptide elongation time for translating the epitope and 35 additional codons. Although the contribution of the peptide elongation time can be corrected (step 6.9.), this assay consequently is most sensitive for measuring initiation kinetics that occur on the order of minutes, which appears to be a generic property of cap-dependent initiation15. Experimentally, it is easy to adapt this assay to study other initiation mechanisms. However, if an initiation mechanism occurs significantly fast on the order of seconds, this assay is unlikely to resolve the initiation kinetics.
The single-molecule approach described here combines single-molecule imaging and extract-based translation to enable measurements of individual cap-dependent translation events in real-time and during active cell-free translation. The assay allows easy transitioning from a bulk translation assay to a single-molecule assay, while preserving the bulk translation kinetics. Thus, single-molecule kinetic observations can be interpreted in direct reference to corresponding bulk results. Previous methods, including recently developed in vivo approaches11,12,13,14, lack the resolution for kinetic measurements of individual translation initiation events and require assumptions of translation mechanisms to extract initiation time averages from experimental observables. The single-molecule method presented here provides the first hypothesis-free approach for quantifying the average initiation time of active cap-dependent translation. Furthermore, although bulk kinetic measurements with a luminescence assay have been utilized to provide the most quantitative and systematic bulk characterizations of cap-dependent translation kinetics to date46,50, the single-molecule assay described here measures average initiation kinetic changes with greater sensitivity than the bulk assay (Figure 7). In addition, the single-molecule data preserves single mRNA translation stochasticity and heterogeneity, properties that are averaged out in bulk measurements. The single-molecule resolution of translation kinetics can be utilized for systematic kinetic analyses to reveal insights of translation mechanisms unattainable with other methods.
This assay is compatible with existing biochemical and genetic approaches that are commonly used for translation studies. For example, the translational function of a specific factor can be studied by generating factor-depleted extract and supplementing it with wild-type or mutant factor. In addition, by supplementing fluorescently labeled factor, studies can potentially investigate the kinetic relationship between factor binding and the progression of translation. With the use of two distinct epitope/antibody pairs, this protocol could potentially be expanded from a one-color assay to a two-color assay that enables simultaneous detection of two distinct coding sequences. This adaptation would allow tracking the translation of different reading frames and could be applied to frameshifting and start site selection studies. Alternatively, a two-color system could be used to detect antibody interactions with tagged polypeptides encoded by upstream and downstream open reading frames to monitor both upstream and downstream translation events on individual mRNAs. These expansions can enable a wide range of mechanistic studies that investigate cap-dependent translation and its regulation.

Translation in eukaryotic systems occurs predominantly through 7-methylguanosine (m7G) cap-dependent pathways1. Studies indicate that the initiation step of eukaryotic translation is rate-limiting and a common target for regulation2,3,4. Mechanisms of cap-dependent translation have been extensively studied using genetic5, biochemical6,7,8, structural9, and genomic10 bulk approaches. Although these methods have identified diverse mechanisms that regulate cap-dependent initiation, their resolution limits them to ensemble averaging of signals from heterogeneous and asynchronous initiation events. More recently, individual in vivo translation events have been visualized by methods that measure fluorescent antibody binding to epitopes on nascent polypeptides11,12,13,14. However, these new approaches are also limited in their ability to resolve individual initiation events because multiple fluorescent antibodies must bind a nascent peptide to allow single translation events to be resolved from a high intracellular fluorescence background. In many biological interactions, resolved individual kinetic events have provided critical insights into understanding complex multistep and repetitive biological processes that are not possible to synchronize at the molecular level. New methods that can track the dynamics of individual translation events are needed for a better understanding of cap-dependent initiation and regulation.
We recently developed an in vitro assay that measures cap-dependent initiation kinetics with single-molecule resolution15. Considering the large number of known and unknown protein factors involved in this initiation pathway3,16, the single-molecule assay was developed to be compatible with existing in vitro cell-free translation systems to benefit from their preservation of cellular factors and robust translation activity17,18,19,20,21,22,23,24,25. Furthermore, the use of cell-free translation systems allows more compatible comparisons between single-molecule observations and previous bulk results. This approach provides a straight-forward integration of new single-molecule kinetic insights into the existing mechanistic framework of cap-dependent initiation. To establish the single-molecule assay, the traditional cell-free translation system is modified in three ways: an epitope-encoding sequence is inserted at the beginning of the open reading frame (ORF) of a reporter mRNA; the 3&#8242; end of the reporter mRNA is biotinylated to facilitate mRNA end-tethering to single-molecule detection surface; and fluorescently-labeled antibodies are supplemented to the translation extract. These modifications require only basic molecular biology techniques and commonly available reagents. Furthermore, these modifications and the single-molecule imaging conditions preserve the translation kinetics of bulk cell-free translation reactions15.
In this assay (Figure 1), 5&#8242;-end capped and 3&#8242;-end biotinylated reporter mRNA is immobilized to a streptavidin-coated detection surface in a flow chamber. The flow chamber is then filled with a cell-free translation mixture supplemented with fluorescently labeled antibodies. After mRNA translation has occurred for approximately 30-40 codons downstream of the epitope sequence26,27, the epitope emerges from the ribosome exit tunnel and becomes accessible to interact with fluorescently-labeled antibody. This interaction is rapid and its detection by single-molecule fluorescence imaging techniques enables tracking of translation kinetics with single-molecule resolution during active cell-free translation. This assay should broadly benefit in vitro studies of cap-dependent translation kinetics and its regulation, particularly for systems with a working bulk in vitro assay.
A prerequisite for establishing this single-molecule assay is a working bulk cell-free translation assay, which can be achieved using translation extract that is either commercially available or prepared following previously described methods28. Eukaryotic translation extract can be obtained from diverse cells, including fungal, mammalian, and plant28. For imaging, this assay requires a TIRF microscope equipped with tunable laser intensity and incident angle, a motorized sample stage, a motorized fluidics system, and sample temperature control device. Such requirements are generic for modern in vitro single-molecule TIRF experiments and may be achieved differently. The experiment presented here uses an objective-type TIRF system made up of commercially available microscope, software, and accessories all listed in the Table of Materials.

<table><tbody><tr><td>100X oil objective, N.A. 1.49</td><td>Olympus</td><td>UAPON 100XOTIRF</td><td></td></tr><tr><td>Acryamide/bis (40%, 19:1)</td><td>Bio-Rad</td><td>161-0144</td><td></td></tr><tr><td>Alkaline liquid detergent</td><td>Decon</td><td>5332</td><td></td></tr><tr><td>Aminosilane (N-(2-Aminoethyl)-3-Aminopropyltrimethoxysilane)</td><td>UCT Specialties, LLC</td><td>A0700</td><td></td></tr><tr><td>Andor ixon Ultra DU 897V EMCCD</td><td>Andor</td><td>DU-897U-CSO-#BV</td><td></td></tr><tr><td>Andor Solis software</td><td>Andor</td><td></td><td>For controlling the Andor EMCCD</td></tr><tr><td>Band-pass filter</td><td>Chroma</td><td>532/640/25</td><td></td></tr><tr><td>Band-pass filter</td><td>Chroma</td><td>NF03-405/488/532/635E-25</td><td></td></tr><tr><td>Biotin-PEG-SVA</td><td>Laysan Bio Inc</td><td>Biotin-PEG-SVA</td><td></td></tr><tr><td>Coenzyme A free acid</td><td>Prolume</td><td>309-250</td><td></td></tr><tr><td>Coolterm software</td><td></td><td></td><td>For controlling the syringe pump</td></tr><tr><td>Desktop computer</td><td>Dell</td><td></td><td>For controlling the microscope, camera, stage, and pump.</td></tr><tr><td>Dichroic mirror</td><td>Semrock</td><td>R405/488/532/635</td><td></td></tr><tr><td>Direct-zol RNA microprep 50RNX</td><td>Fisher Scientific</td><td>NC1139450</td><td></td></tr><tr><td>Dual-Luciferase Reporter Assay System</td><td>Promega</td><td>E1910</td><td></td></tr><tr><td>Epoxy</td><td>Devcon</td><td>14250</td><td></td></tr><tr><td>Firefly luciferin D-Luciferin free acid</td><td>Prolume</td><td>306-250</td><td></td></tr><tr><td>Glacial acetic acid</td><td>Fisher Scientific</td><td>BP1185500</td><td></td></tr><tr><td>Hydrogen perioxide</td><td>Sigma-Aldrich</td><td>216763-500ML</td><td></td></tr><tr><td>Immersion oil</td><td>Olympus</td><td>Z-81226A</td><td>Low auto-fluorescence</td></tr><tr><td>Luciferase Assay System</td><td>Promega</td><td>E1500</td><td></td></tr><tr><td>MEGASCRIPT T7 Transcription Kit</td><td>Thermo fisher</td><td>AM1334</td><td></td></tr><tr><td>Methanol</td><td>Fisher Scientific</td><td>MMX04751</td><td></td></tr><tr><td>Microscope</td><td>Olympus</td><td>IX83</td><td></td></tr><tr><td>Microscope slide</td><td>Thermo Scientific</td><td>3048</td><td></td></tr><tr><td>Monoclonal anti-FLAG M2-Cy3</td><td>Sigma-Aldrich</td><td>A9594</td><td></td></tr><tr><td>mPEG-SVA</td><td>Laysan Bio Inc</td><td>mPEG-SVA-5000</td><td></td></tr><tr><td>MS(PEG)4</td><td>Thermo Scientific</td><td>22341</td><td></td></tr><tr><td>NaCl (5M)</td><td>Thermo Scientific</td><td>AM9760G</td><td></td></tr><tr><td>No 1.5 microscope Cover glass</td><td>Fisherband</td><td>12-544-C</td><td></td></tr><tr><td>Olympus Laser, 532nm 100mM</td><td>Olympus digital Laser system</td><td>CMR-LAS 532nm 100mW</td><td></td></tr><tr><td>Olympus TirfCtrl software</td><td>Olympus</td><td></td><td>For controlling the laser intensity and incident angle</td></tr><tr><td>Optical table</td><td>TMC vibration control</td><td>63-563</td><td>With vibration isolation</td></tr><tr><td>Phenol chloroform isoamyl alcohol mix</td><td>Sigma-Aldrich</td><td>77617-100ml</td><td></td></tr><tr><td>Pierce RNA 3&#039; End Biotinylation Kit</td><td>Thermo Scientific</td><td>20160</td><td></td></tr><tr><td>Potassium hydroxide pellets</td><td>Sigma-Aldrich</td><td>P1767-500G</td><td></td></tr><tr><td>Prior motorized XY translation stage</td><td>Prior</td><td>PS3J100</td><td></td></tr><tr><td>Prior PriorTest software</td><td>Prior</td><td></td><td>For controlling the Prior motorized stage</td></tr><tr><td>Recombinant RNasin RNase Inhibitor</td><td>Promega</td><td>N2515</td><td></td></tr><tr><td>Stage top Incubator</td><td>In vivo scientific (world precision Instruments)</td><td>98710-1</td><td>With a custom built acrylic cage</td></tr><tr><td>Staining jar</td><td>Fisher Scientific</td><td>08-817</td><td></td></tr><tr><td>Streptavidin</td><td>Thermo Scientific</td><td>43-4301</td><td></td></tr><tr><td>Sulfuric acid</td><td>Fisher Scientific</td><td>A300212</td><td></td></tr><tr><td>SYBR green II</td><td>Fisher Scientific</td><td>S7564</td><td></td></tr><tr><td>Syringe</td><td>Hamilton</td><td>1725RN</td><td></td></tr><tr><td>Syringe pump</td><td>Harvard apparatus</td><td>55-3333</td><td></td></tr><tr><td>Tris (1M), pH = 7.0</td><td>Thermo Scientific</td><td>AM9850G</td><td></td></tr><tr><td>Ultrasonic Bath</td><td>Branson</td><td>CPX1800H</td><td></td></tr><tr><td>Urea</td><td>Sigma-Aldrich</td><td>U5378-500G</td><td></td></tr><tr><td>Vaccinia Capping system</td><td>New England Biolabs</td><td>M2080S</td><td></td></tr><tr><td>Zymo-Spin IC Columns</td><td>Zymo Research</td><td>C1004</td><td></td></tr></tbody></table>

an in vitro single-molecule imaging assay for the analysis of cap-dependent translation kinetics

Cap-dependent protein synthesis is the predominant translation pathway in eukaryotic cells. While various biochemical and genetic approaches have allowed extensive studies of cap-dependent translation and its regulation, high resolution kinetic characterization of this translation pathway is still lacking. Recently, we developed an in vitro assay to measure cap-dependent translation kinetics with single-molecule resolution. The assay is based on fluorescently labeled antibody binding to nascent epitope-tagged polypeptide. By imaging the binding and dissociation of antibodies to and from nascent peptide&#8211;ribosome&#8211;mRNA complexes, the translation progression on individual mRNAs can be tracked. Here, we present a protocol for establishing this assay, including mRNA and PEGylated slide preparations, real-time imaging of translation, and analysis of single molecule trajectories. This assay enables tracking of individual cap-dependent translation events and resolves key translation kinetics, such as initiation and elongation rates. The assay can be widely applied to distinct translation systems and should broadly benefit in vitro studies of cap-dependent translation kinetics and translational control mechanisms.

Following the protocol described enables the imaging of individual antibody interactions with nascent N-terminal-tagged polypeptides with single-molecule resolution during active cell-free translation of 3&#39; end-tethered reporter mRNA (Figure 1). A minimal demonstration experiment is reported with the use of three synthetic mRNAs: LUC (encoding untagged luciferase), LUCFLAG (encoding 3xFLAG-tagged luciferase), and hp-LUCFLAG (LUCFLAG with a stable stem-loop in the 5&#39; leader region) (Figure 2C). The use of luciferase-encoding mRNA enables comparisons between single-molecule observations and bulk luminescence measurements. Integrity of 3&#39; biotinylated mRNA was assessed by denaturing polyacrylamide gel electrophoresis (PAGE) and RNA staining. Good quality mRNA shows a single clean band and should not show additional faster migrating signals (Figure 3A). Saccharomyces cerevisiae translation extract was used here and prepared following a protocol that was previously described30 and modified15. Bulk measurements of luciferase activity in cell-free translation reactions with the yeast extract and biotinylated LUCFLAG mRNA that either lacks or contains an m7G cap shows cap stimulation of LUCFLAG mRNA translation (Figure 3B).
For single-molecule imaging, translation extract was supplemented with 67 nM of Cy3-labeled anti-FLAG antibody (Cy3-&#945;FLAG). This antibody had a high labeling ratio of 2-7 dyes per antibody. The 532 nm laser was set to 10 &#956;W at the objective. The laser incident angle was set to 71.5&#730;, which was greater than the critical angle of 63.8&#730; for our set up. A low level of fluorescence was imaged from detection surfaces that contain mRNA but lack translation mix (Figure 4A). For channel dimensions of approximately 2 mm (width) x 50 &#956;m (depth) x 24 mm (length), a flow delivery rate of 150 &#956;l/min yielded a reagent exchange time of 3 &#8211; 4 s. Within 5 s of translation mix delivery, the background fluorescence level increases due to the diffusing fluorescent antibodies. Approximately 2 min after translation mix delivery to LUCFLAG mRNA-containing channels, Cy3 spots began to appear in a field of view and continued to accumulate for ~30 min (Figure 4B). Nonspecific antibody binding to surface, measured using the 3xFLAG-lacking LUC mRNA (Figure 2C), remained at a very low level15. The signals from mRNA/ribosome/peptide-bound Cy3- &#945;FLAG were clearly visible with the optimized laser incident angle but could be masked by a high fluorescence background with lower laser incident angles (Figure 4C).
To analyze Cy3-&#945;FLAG binding events, the fluorescence intensities of selected Cy3-&#945;FLAG spots (Figure 5A) were calculated frame by frame for the entire movie and then connected to form an intensity trajectory (Figure 5B). Raw trajectories can appear noisy and may require refinement with a non-linear forward-backward filter to reduce background noise41. Further refinement could be achieved using step-detection algorithms to digitize trajectories42. For all trajectories, a universal background fluorescence level increase appears during reagent exchange due to the diffusing fluorescent antibodies in the translation mix (Figure 5B). Antibody binding to a nascent polypeptide caused an instantaneous increase in fluorescence whereas antibody/polypeptide dissociation from mRNA causes an instantaneous fluorescence decrease (Figure 5B).
The dwell time of antibody binding can be used to measure the total decoding time of an open reading frame. The dwell time histogram for LUCFLAG mRNA fits to a log-normal distribution (Figure 6A) and yielded a peptide synthesis rate of 2.5 &#177; 0.1 amino acids per second15. The first arrival time histogram fits to a shifted (3-parameter) log-normal function (Figure 6B). The wide spread of the first arrival time histogram shows the high degree of heterogeneity in single mRNA translation activity. The histogram indicated that the first peptide synthesis event on single LUCFLAG mRNA molecules could begin as early as 2 min, and as late as 20 min, after translation mix delivery (Figure 6B). Compared to translation with LUCFLAG mRNA, the first arrival time histogram with hp-LUCFLAG mRNA translation showed slower antibody binding (Figure 7A). Using bulk luminescence measurements from cell-free translation reactions with these same mRNAs, however, does not resolve differences in translation kinetics (Figure 7B,C). These results demonstrate the higher resolution of our method compared to bulk kinetic luciferase activity measurements for detecting small changes in initiation kinetics.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61648/61648fig01.jpg" /> 
Figure 1: Protocol flow chart.&#160;Schematic representations of coverslip and slide preparation, single-molecule chamber assembly, TIRF imaging and data acquisition, and data analysis steps are shown. The TIRF imaging step includes schematic depictions of mRNA immobilization and translation in a flow channel. Detection surface components, fluorescently labeled antibody, and cell extract components are indicated. <a href="https://www.jove.com/files/ftp_upload/61648/61648fig01large.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/61648/61648fig2v2.jpg" /> 
Figure 2: mRNA designs for the single-molecule assay.&#160;(A) To adapt a bulk translation assay to the single-molecule assay, the DNA template of the bulk reporter ORF was modified with an N-terminal epitope tag sequence insertion to generate a tagged reporter-encoding ORF. ORF and N-terminal tag-coding regions are indicated. (B) mRNAs were 5&#8242;-m7G capped to allow cap-dependent translation and 3&#8242;-biotinylated for their immobilization to single-molecule detection surface. 5&#8242;-m7G cap and 3&#8242;-biotin are indicated on untagged and tagged ORF RNAs. (C) Luciferase-encoding mRNAs were used to demonstrate the single-molecule assay. LUC and LUCFLAG mRNAs encode luciferase that lacks and contains an N-terminal 3xFLAG tag, respectively. hp-LUCFLAG mRNA contains an additional insertion that introduces a hairpin secondary structure in the LUCFLAG mRNA 5&#8242;-leader. The hairpin thermostability, 3&#8242;-poly(A) tail, and 3&#8242;-biotin are indicated. All three mRNAs included a 30 nt 3&#8242;-poly(A) tail for more efficient cap-dependent translation. <a href="https://www.jove.com/files/ftp_upload/61648/61648fig02large.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/61648/61648fig3v2.jpg" /> 
Figure 3: Single-molecule reporter mRNA integrity and bulk translation.&#160;(A) Representative denaturing PAGE imaging of single-molecule reporter mRNA. 5&#8242;-m7G capped and 3&#8242;-biotinylated LUCFLAG mRNA was loaded onto a denaturing 10% polyacrylamide gel next to an RNA ladder. (B) Single-molecule reporter mRNA preserved 5&#39; cap-dependence in bulk cell-free translation reactions. 3&#8242;-biotinylated LUCFLAG mRNA that either lacked (&#8211;cap) or contained (+ cap) a 5&#8242;-m7G cap was translated in S. cerevisiae translation extract for 30 min at 25 &#730;C. Luciferase activity was measured from two independent reactions and normalized to the activity with &#8211;cap mRNA, which is arbitrarily set to 1.0. Standard deviations from mean values are indicated by error bars. <a href="https://www.jove.com/files/ftp_upload/61648/61648fig03large.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/61648/61648fig04.jpg" /> 
Figure 4: Imaging of detection surfaces containing immobilized mRNA.&#160;(A) Background fluorescence in the absence of translation mix. (B) Cy3 fluorescent spots in a field of view 30 min after the translation mix delivery and with an optimized laser incident angle. (C) Imaged field of view 30 min after translation mix delivery and with a low laser incident angle. <a href="https://www.jove.com/files/ftp_upload/61648/61648fig04large.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/61648/61648fig05.jpg" /> 
Figure 5: Image analysis.&#160;(A) Cy3 spots in a field of view without (left panel) or with (right panel) spot selection. (B) Example single-molecule trajectory for a selected Cy3 spot. The black arrow indicates the rise in background fluorescence due to diffusing antibody during translation mix delivery. The green arrow indicates the sudden increase in fluorescence intensity due to Cy3-&#945;FLAG binding. The red arrow indicates the sudden decrease in fluorescence intensity due to Cy3-&#945;FLAG/nascent peptide dissociation from mRNA. The dwell time of a Cy3-&#945;FLAG binding event is indicated with a double-headed arrow. Raw, filtered, and digitized data are indicated. <a href="https://www.jove.com/files/ftp_upload/61648/61648fig05large.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/61648/61648fig06.jpg" /> 
Figure 6: Kinetic analysis of Cy3-&#945;FLAG binding with LUCFLAG mRNA translation.&#160;(A) The Cy3-&#945;FLAG binding dwell time histogram fits to a log-normal distribution. (B) The Cy3-&#945;FLAG binding first arrival time histogram fits to a shifted (3-parameter) log-normal function. Adapted from Wang et al.15 with permission. <a href="https://www.jove.com/files/ftp_upload/61648/61648fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/61648/61648fig7v3.jpg" /> 
Figure 7: The single-molecule assay has a higher resolution for initiation kinetics than the bulk luminescence assay.&#160;(A) First arrival time histograms for Cy3-&#945;FLAG binding with translation of LUCFLAG and hp-LUCFLAG mRNAs revealed slower initiation caused by a small hairpin structure in the mRNA 5&#8242;-leader. N = 10650 trajectories (LUCFLAG), combined from 3 data sets and N = 4079 trajectories (hp-LUCFLAG), combined from 2 data sets. (B,C) Bulk luciferase activity assay kinetic measurements did not resolve differences in initiation kinetics for LUCFLAG (N = 9 data sets) and hp-LUCFLAG (N = 6 data sets) mRNA translation. (B) Bulk luminescence kinetics. (C) Scatter plots of first round translation times determined by Gaussian fitting to the second derivative of the luminescence kinetics curves in (B), as described by Vassilenko et al.46. The red circles and bars in (C) represent the mean value and standard deviation of the calculated first round translation time, respectively. Adapted from Wang et al.15 with permission. <a href="https://www.jove.com/files/ftp_upload/61648/61648fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about An In Vitro Single-Molecule Imaging Assay for the Analysis of Cap-Dependent Translation Kinetics at JoVE.com

An In Vitro Single-Molecule Imaging Assay for the Analysis of Cap-Dependent Translation Kinetics

Tracking individual translation events allows for high-resolution kinetic studies of cap-dependent translation mechanisms. Here we demonstrate an in vitro single-molecule assay based on imaging interactions between fluorescently labeled antibodies and epitope-tagged nascent peptides. This method enables single-molecule characterization of initiation and peptide elongation kinetics during active in vitro cap-dependent translation.

An In Vitro Single-Molecule Imaging Assay for the Analysis of Cap-Dependent Translation Kinetics

Cap-dependent protein synthesis is the predominant translation pathway in eukaryotic cells. While various biochemical and genetic approaches have allowed ...

an-vitro-single-molecule-imaging-assay-for-analysis-cap-dependent

Nanyang Technological University

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JoVE Journal

Biochemistry

3.0K Views.  Memorial Sloan Kettering Cancer Center. Het volgen van individuele vertaalgebeurtenissen maakt kinetische studies met hoge resolutie van cap-afhankelijke vertaalmechanismen mogelijk. Hier demonstreren we een in vitro single-molecule assay op basis van beeldvormende interacties tussen fluorescerend gelabelde antilichamen en epitoop-gelabelde ontluikende peptiden. Deze methode maakt single-molecule karakterisering van initiatie en peptide rekkinetiek mogelijk tijdens actieve in vitro cap-afhanke...

Video: An In Vitro Single-Molecule Imaging Assay for the Analysis of Cap-Dependent Translation Kinetics

Quantitative Immunofluorescence to Measure Global Localized Translation

The mechanisms regulating mRNA translation are involved in various biological processes, such as germ line development, cell differentiation, and organogenesis, as well as in multiple diseases. Numerous publications have convincingly shown that specific mechanisms tightly regulate mRNA translation. Increased interest in the translation-induced regulation of protein expression has led to the development of novel methods to study and follow de novo protein synthesis in cellulo. However, most of these methods are complex, making them costly and often limiting the number of mRNA targets that can be studied. This manuscript proposes a method that requires only basic reagents and a confocal fluorescence imaging system to measure and visualize the changes in mRNA translation that occur in any cell line under various conditions. This method was recently used to show localized translation in the subcellular structures of adherent cells over a short period of time, thus offering the possibility of visualizing de novo translation for a short period during a variety of biological processes or of validating changes in translational activity in response to specific stimuli.

This manuscript describes a method to visualize and quantify localized translation events in subcellular compartments. The approach proposed in this manuscript requires a basic confocal imaging system and reagents and is rapid and cost-effective.

The mechanisms regulating mRNA translation are involved in various biological processes, such as germ line development, cell differentiation, and ...

Single-Molecule Analysis of Sf9 Purified Superprocessive Kinesin-3 Family Motors

A complex cellular environment poses challenges for single-molecule motility analysis. However, advancement in imaging techniques have improved single-molecule studies and has gained immense popularity in detecting and understanding the dynamic behavior of fluorescent-tagged molecules. Here, we describe a detailed method for&#160;in vitro single-molecule studies of kinesin-3 family motors using Total Internal Reflection Fluorescence (TIRF) microscopy. Kinesin-3 is a large family that plays critical roles in cellular and physiological functions ranging from intracellular cargo transport to cell division to development. We have shown previously that constitutively active dimeric kinesin-3 motors exhibit fast and superprocessive motility with high microtubule affinity at the single-molecule level using cell lysates prepared by expressing motor in mammalian cells. Our lab studies kinesin-3 motors and their regulatory mechanisms using cellular, biochemical and biophysical approaches, and such studies demand purified proteins at a large scale. Expression and purification of these motors using mammalian cells would be expensive and time-consuming, whereas expression in a prokaryotic expression system resulted in significantly aggregated and&#160;inactive protein. To overcome the limitations posed by bacterial purification systems and mammalian cell lysate, we have established a robust Sf9-baculovirus expression system to express and purify these motors. The kinesin-3 motors are C-terminally tagged with 3-tandem fluorescent proteins (3xmCitirine or 3xmCit) that provide enhanced signals and decreased photobleaching. In vitro single-molecule and multi-motor gliding analysis of Sf9 purified proteins demonstrate that kinesin-3 motors are fast and superprocessive akin to our previous studies using mammalian cell lysates. Other applications using these assays include detailed knowledge of oligomer conditions of motors, specific binding partners paralleling biochemical studies, and their kinetic state.

This study details purification of KIF1A(1-393LZ), a member of kinesin-3 family, using Sf9-baculovirus expression system. In vitro single-molecule and multi-motor gliding analysis of these purified motors exhibited robust motility properties comparable to motors from mammalian cell lysate. Thus, Sf9-baculovirus system is amenable to express and purify motor protein of interest.

A complex cellular environment poses challenges for single-molecule motility analysis. However, advancement in imaging techniques have improved ...

Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence (TIRF) Microscopy

Traditionally, the actin and microtubule cytoskeletons have been studied as separate entities, restricted to specific cellular regions or processes, and regulated by different suites of binding proteins unique for each polymer. Many studies now demonstrate that the dynamics of both cytoskeletal polymers are intertwined and that this crosstalk is required for most cellular behaviors. A number of proteins involved in actin-microtubule interactions have already been identified (i.e., Tau, MACF, GAS, formins, and more) and are well characterized with regard to either actin or microtubules alone. However, relatively few studies showed assays of actin-microtubule coordination with dynamic versions of both polymers. This may occlude emergent linking mechanisms between actin and microtubules. Here, a total internal reflection fluorescence (TIRF) microscopy-based in vitro reconstitution technique permits the visualization of both actin and microtubule dynamics from the one biochemical reaction. This technique preserves the polymerization dynamics of either actin filament or microtubules individually or in the presence of the other polymer. Commercially available Tau protein is used to demonstrate how actin-microtubule behaviors change in the presence of a classic cytoskeletal crosslinking protein. This method can provide reliable functional and mechanistic insights into how individual regulatory proteins coordinate actin-microtubule dynamics at a resolution of single filaments or higher-order complexes.

Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence TIRF Microscopy

This protocol is a guide for visualizing dynamic actin and microtubules using an in vitro total internal fluorescence (TIRF) microscopy assay.

Traditionally, the actin and microtubule cytoskeletons have been studied as separate entities, restricted to specific cellular regions or processes, and ...

Determination of In Vitro and Cellular Turn-on Kinetics for Fluorogenic RNA Aptamers

Fluorogenic RNA aptamers have been applied in live cells to tag and visualize RNAs, report on gene expression, and activate fluorescent biosensors that detect levels of metabolites and signaling molecules. In order to study dynamic changes in each of these systems, it is desirable to obtain real-time measurements, but the accuracy of the measurements depends on the kinetics of the fluorogenic reaction being faster than the sampling frequency. Here, we describe methods to determine the in vitro and cellular turn-on kinetics for fluorogenic RNA aptamers using a plate reader equipped with a sample injector and a flow cytometer, respectively. We show that the in vitro kinetics for the fluorescence activation of the Spinach2 and Broccoli aptamers can be modeled as two-phase association reactions and have differing fast phase rate constants of 0.56 s&#8722;1 and 0.35 s&#8722;1, respectively. In addition, we show that the cellular kinetics for the fluorescence activation of Spinach2 in Escherichia coli, which is further limited by dye diffusion into the Gram-negative bacteria, is still sufficiently rapid to enable accurate sampling frequency on the minute timescale. These methods to analyze fluorescence activation kinetics are applicable to other fluorogenic RNA aptamers that have been developed.

Determination of In Vitro and Cellular Turn-on Kinetics for Fluorogenic RNA Aptamers

The protocol presents two methods to determine the kinetics of the fluorogenic RNA aptamers Spinach2 and Broccoli. The first method describes how to measure fluorogenic aptamer kinetics in vitro with a plate reader, while the second method details the measurement of fluorogenic aptamer kinetics in cells by flow cytometry.

Fluorogenic RNA aptamers have been applied in live cells to tag and visualize RNAs, report on gene expression, and activate fluorescent biosensors that ...

In Vitro Transcribed RNA-based Luciferase Reporter Assay to Study Translation Regulation in Poxvirus-infected Cells

Every poxvirus mRNA transcribed after viral DNA replication has an evolutionarily conserved, non-templated 5&#39;-poly(A) leader in the 5&#39;-UTR. To dissect the role of 5&#39;-poly(A) leader in mRNA translation during poxvirus infection we developed an in vitro transcribed RNA-based luciferase reporter assay. This reporter assay comprises of four core steps: (1) PCR to amplify the DNA template for in vitro transcription; (2) in vitro transcription to generate mRNA using T7 RNA polymerase; (3) Transfection to introduce in vitro transcribed mRNA into cells; (4) Detection of luciferase activity as the indicator of translation. The RNA-based luciferase reporter assay described here circumvents issues of plasmid replication in poxvirus-infected cells and cryptic transcription from the plasmid. This protocol can be used to determine translation regulation by cis-elements in an mRNA including 5&#39;-UTR and 3&#39;-UTR in systems other than poxvirus-infected cells. Moreover, different modes of translation initiation like cap-dependent, cap-independent, re-initiation, and internal initiation can be investigated using this method.

We present a protocol to study mRNA translation regulation in poxvirus-infected cells using in vitro Transcribed RNA-based luciferase reporter assay. The assay can be used for studying translation regulation by cis-elements of an mRNA, including 5&#8217;-untranslated region (UTR) and 3&#8217;-UTR. Different translation initiation modes can also be examined using this method.

Every poxvirus mRNA transcribed after viral DNA replication has an evolutionarily conserved, non-templated 5&#39;-poly(A) leader in the 5&#39;-UTR. To ...

Immunology and Infection

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 ...

Biology

Real-time Imaging of Single Engineered RNA Transcripts in Living Cells Using Ratiometric Bimolecular Beacons

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the development of diverse techniques to visualize individual RNA transcripts in single living cells. One promising technique that has recently been described utilizes an oligonucleotide-based optical probe, ratiometric bimolecular beacon (RBMB), to detect RNA transcripts that were engineered to contain at least four tandem repeats of the RBMB target sequence in the 3&#8217;-untranslated region. RBMBs are specifically designed to emit a bright fluorescent signal upon hybridization to complementary RNA, but otherwise remain quenched. The use of a synthetic probe in this approach allows photostable, red-shifted, and highly emissive organic dyes to be used for imaging. Binding of multiple RBMBs to the engineered RNA transcripts results in discrete fluorescence spots when viewed under a wide-field fluorescent microscope. Consequently, the movement of individual RNA transcripts can be readily visualized in real-time by taking a time series of fluorescent images. Here we describe the preparation and purification of RBMBs, delivery into cells by microporation and live-cell imaging of single RNA transcripts.

Ratiometric bimolecular beacons (RBMBs) can be used to image single engineered RNA transcripts in living cells. Here, we describe the preparation and purification of RBMBs, delivery of RBMBs into cells by microporation and fluorescent imaging of single RNA transcripts in real-time.

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the ...

Micromanipulation Techniques Allowing Analysis of Morphogenetic Dynamics and Turnover of Cytoskeletal Regulators

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how fluorescent recovery after photobleaching (FRAP) and photoactivation techniques can be used to study the spatiotemporal dynamics of proteins in subcellular locations. We also show how these techniques enable straightforward determination of various parameters linked to actin cytoskeletal regulation and cell motility. Moreover, the microinjection of cells is additionally described as an alternative treatment (potentially preceding or complementing the aforementioned photomanipulation techniques) to trigger instantaneous effects of translocated proteins on cell morphology and function. Micromanipulation such as protein injection or local application of plasma membrane-permeable drugs or cytoskeletal inhibitors can serve as powerful tool to record immediate consequences of a given treatment on cell behavior at the single cell and subcellular level. This is exemplified here by immediate induction of lamellipodial cell edge protrusion by the injection of recombinant Rac1 protein, as established a quarter-century ago. In addition, we provide a protocol for determining the turnover of enhanced green fluorescent protein (EGFP)-VASP, an actin filament polymerase prominently accumulating at lamellipodial tips of B16-F1 cells, employing FRAP and including associated data analysis and curve fitting. We also present guidelines for estimating the rates of lamellipodial actin network polymerization, as exemplified by cells expressing EGFP-tagged &#946;-actin. Finally, instructions are given for how to investigate the rates of actin monomer mobility within the cell cytoplasm, followed by actin incorporation at sites of rapid filament assembly, such as the tips of protruding lamellipodia, using photoactivation approaches. None of these protocols is&#160;restricted to components or regulators of the actin cytoskeleton, but can easily be extended to explore in analogous fashion the spatiotemporal dynamics and function of proteins in various different subcellular structures or functional contexts.

We describe how micro- and photomanipulation techniques such as FRAP and photoactivation enable the determination of motility parameters and the spatiotemporal dynamics of proteins within migrating cells. Experimental readouts include subcellular dynamics and turnover of motility regulators or of the underlying actin cytoskeleton.

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how ...

Probing mRNA Kinetics in Space and Time in Escherichia coli using Two-Color Single-Molecule Fluorescence In Situ Hybridization

Single-molecule fluorescence in situ hybridization (smFISH) allows for counting the absolute number of mRNAs in individual cells. Here, we describe an application of smFISH to measure the rates of transcription and mRNA degradation in Escherichia coli. As smFISH is based on fixed cells, we perform smFISH at multiple time points during a time-course experiment, i.e., when cells are undergoing synchronized changes upon induction or repression of gene expression. At each time point, sub-regions of an mRNA are spectrally distinguished to probe transcription elongation and premature termination. The outcome of this protocol also allows for analyzing intracellular localization of mRNAs and heterogeneity in mRNA copy numbers among cells. Using this protocol many samples (~50) can be processed within 8 h, like the amount of time needed for just a few samples. We discuss how to apply this protocol to study the transcription and degradation kinetics of different mRNAs in bacterial cells.

Probing mRNA Kinetics in Space and Time in Escherichia coli using Two-Color Single-Molecule Fluorescence In Situ Hybridization

This protocol describes an application of single-molecule fluorescence in situ hybridization (smFISH) to measure the in vivo kinetics of mRNA synthesis and degradation.

Single-molecule fluorescence in situ hybridization (smFISH) allows for counting the absolute number of mRNAs in individual cells. Here, we describe an ...

Live-cell Imaging of Single-Cell Arrays (LISCA) - a Versatile Technique to Quantify Cellular Kinetics

Live-cell Imaging of Single-Cell Arrays (LISCA) is a versatile method to collect time courses of fluorescence signals from individual cells in high throughput. In general, the acquisition of single-cell time courses from cultured cells is hampered by cell motility and diversity of cell shapes. Adhesive micro-arrays standardize single-cell conditions and facilitate image analysis. LISCA combines single-cell microarrays with scanning time-lapse microscopy and automated image processing. Here, we describe the experimental steps of taking single-cell fluorescence time courses in a LISCA format. We transfect cells adherent to a micropatterned array using mRNA encoding for enhanced green fluorescent protein (eGFP) and monitor the eGFP expression kinetics of hundreds of cells in parallel via scanning time-lapse microscopy. The image data stacks are automatically processed by newly developed software that integrates fluorescence intensity over selected cell contours to generate single-cell fluorescence time courses. We demonstrate that eGFP expression time courses after mRNA transfection are well described by a simple kinetic translation model that reveals expression and degradation rates of mRNA. Further applications of LISCA for event time correlations of multiple markers in the context of signaling apoptosis are discussed.

Live-cell Imaging of Single-Cell Arrays LISCA - a Versatile Technique to Quantify Cellular Kinetics

We present a method for the acquisition of fluorescence reporter time courses from single cells using micropatterned arrays. The protocol describes the preparation of single-cell arrays, the setup and operation of live-cell scanning time-lapse microscopy and an open-source image analysis tool for automated preselection, visual control and tracking of cell-integrated fluorescence time courses per adhesion site.

Live-cell Imaging of Single-Cell Arrays (LISCA) is a versatile method to collect time courses of fluorescence signals from individual cells in high ...

An In Vitro Single-Molecule Imaging Assay for the Analysis of Cap-Dependent Translation Kinetics

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