The authors thank Marit Warrer for excellent technical assistance. This work was supported by the Danish Council for Independent Research | Natural Sciences [1323-00343B] and the Danish National Research Foundation [DNRF120].

1. Preparation of bacterial cultures.
<ol>
	<li>Grow all cultures in Erlenmeyer flasks with a culture/flask volume ratio of at most 1/6. In a typical experimental setup, grow the cultures at 37.0 &#176;C and shaking at 160 rpm in morpholinepropanesulfonic acid (MOPS) minimal medium18 supplemented with the preferred carbon source, for example 0.2% glucose, for at least 10 consecutive generations before RNA harvest.</li>
	<li>The day before RNA harvest, dilute an outgrown culture of the desired strain in MOPS minimal medium in such a way that it will reach the desired OD436 at the desired time the following day. Calculate the volume of outgrown culture to use for inoculation by the formula: 
	<img alt="Equation 1" src="/files/ftp_upload/56212/56212eq1.jpg" /> 
	NOTE: Here, ODnew denotes the desired OD436 after incubation, Vnew denotes the volume of the diluted culture, ODold denotes the OD436 of the outgrown culture diluted from, G denotes the number of generations the culture will grow, from the time of dilution (t1) to the time it is needed the day after (t2) and can be calculated as: time difference (in min) between t1 and t2 divided by the generation time (in min). To ensure steady-state growth at the time of sampling, G should be &#8805;10; Vold denotes the volume of the out-grown culture that should be transferred to the fresh medium. This equation is an approximation that does not account for any lag phase; a phenomenon often observed after dilution of an out-grown culture in fresh media.
	<ol>
		<li>Growth conditions will vary depending on the purpose of the experiment, but to limit cell-to-cell variation in tRNA levels and to increase the reproducibility of the results, (recommended) grow the cultures to steady state prior to sampling19.</li>
	</ol>
	</li>
</ol>
2. Preparation of spike-in cells
NOTE: The E. coli strain MAS1074, which expresses the selC gene encoding tRNAselC under the control of an isopropyl &#946;-D-1-thiogalactopyranoside (IPTG)-inducible promoter is used as the spike-in strain.
<ol>
	<li>Dilute an outgrown culture of MAS1074 to an OD436 of 0.05 in MOPS minimal medium supplemented with 100 &#181;g/mL ampicillin and 0.2% glucose. Grow the culture at 37 &#176;C shaking at 160 rpm. 
	NOTE: See the Discussion for the use of an alternative reference strain.</li>
	<li>At OD436 ~0.1, add IPTG to a final concentration of 1 mM to induce the expression of tRNAselC. Grow the culture to OD436 ~0.5 (~3-4 h of growth) and move the culture flask to an ice bath. Keep the spike-in cell culture on ice until all samples for RNA purification have been harvested.</li>
</ol>
3. Harvest of experimental samples.
<ol>
	<li>Pre-warm (37 &#176;C) one capped 100 mL Erlenmeyer flask (or similar) containing 4 mL 10 % (w/v) trichloroacetic acid (TCA) for each sample by placing it in a heated water bath. 
	NOTE: CAUTION, TCA decomposes upon heating, producing toxic and corrosive fumes including hydrogen chloride and chloroform. The solution in water is a strong acid; it reacts violently with bases and is corrosive to many metals. Proper precaution should be taken when using this liquid.</li>
	<li>Immediately prior to cell harvest, measure and record the OD436 of the culture.</li>
	<li>To harvest the cells, transfer 4 mL of culture to a pre-warmed flask containing TCA.</li>
	<li>Mix the sample by swirling the flask by hand for 5 s. Mixing with TCA immediately disables all enzymatic activities and thus preserves the charging levels of the tRNAs. 
	NOTE: If multiple samples are to be collected, keep harvested samples in TCA on ice until all samples have been collected.</li>
</ol>
4. Addition of spike-in cells
NOTE: Preparation of spike-in cells is described in Step 2.
<ol>
	<li>When all samples for RNA preparation have been collected, add 5% spike-in cells (based on OD) to each sample. 
	NOTE: The volume of spike-in cell culture needed is calculated by the formula: 
	<img alt="Equation 2" src="/files/ftp_upload/56212/56212eq2.jpg" /> 
	where Vspike denotes the volume of spike-in cell culture needed, Vexp denotes the volume of harvested experimental cell culture (without TCA), ODexp denotes the OD436 of the experimental cell culture at the time of harvest into TCA. ODspike denotes the OD436 of the spike-in cell culture. If multiple samples are harvested at different ODs, the volume of spike-in cells added should be adjusted according to the formula above for each sample.</li>
	<li>To quantify the contribution from the spike-in cells to the RNA species of interest, prepare a sample only containing spike-in cells by mixing 4 mL spike-in cells with 4 mL TCA. Prepare RNA from this sample in the same way as for the other samples.</li>
	<li>Optionally, take another flask containing only the experimental sample, without spike-in cells, and include it to quantify the contribution of the experimental sample to the tRNAselC levels; the endogenous levels of tRNAselC in E. coli K-12 are negligibly low.</li>
</ol>
5. Preparation of RNA
NOTE: The RNA preparation protocol described here is essentially that described by Varshney et al.1; To avoid deacylation of the aminoacylated tRNAs during purification it is important to keep the samples at acidic pH and at 0 &#176;C throughout the course of purification. Use clean sterile tubes/flasks to hold the samples and solutions, and prepare all solutions with ultrapure (18.2 M&#937;&#183;cm resistivity) water.
<ol>
	<li>Pour 8 mL of each sample into an ice-chilled centrifuge tube. Centrifuge samples for 10 min at 9,500 x g at 4 &#176;C. Completely remove supernatant and resuspend in 0.3 mL cold 0.3 M sodium acetate, pH 4.5 and 10 mM EDTA.</li>
	<li>Transfer each sample to a cold 1.5 mL microcentrifuge tube and add 0.3 mL phenol equilibrated with the same buffer. 
	NOTE: CAUTION, phenol produces toxic fumes and is highly corrosive to the skin.</li>
	<li>Vortex each sample 10x for 15 s with at least 1 min between each vortex. In between vortexing keep the samples at 0 &#176;C. Use the frequent pauses to ensure that the samples remain cold.</li>
	<li>Centrifuge samples for 15 min at 20,000 x g at 4 &#176;C. Transfer the water phase (upper phase) to new 1.5 mL microcentrifuge tubes. Add 0.3 mL cold phenol and vortex 4x 15 s as before.</li>
	<li>Centrifuge for 10 min at 20,000 x g at 4 &#176;C. Transfer the water phase to new, cold 1.5 mL microcentrifuge tubes containing 2.5 times the volume of 96% ethanol (~750 &#181;L). Precipitate the RNA by incubating the tubes for 1 h at -20 &#176;C or overnight at -80 &#176;C.</li>
	<li>Centrifuge samples for 30 min at 20,000 x g at 4 &#176;C. Discard the supernatant and carefully add 1 mL cold 70% ethanol without disturbing the RNA pellet. Gently invert once to wash the insides of the tubes with ethanol. 
	NOTE: The pellet can be hard to see at this step.</li>
	<li>Centrifuge for 10 min at 20,000 x g at 4 &#176;C. Completely remove the supernatant and air-dry the pellet until no ethanol is left. Resuspend the pellet by vigorously vortexing in 20 &#181;L cold 10 mM sodium acetate pH 4.5 and 1 mM EDTA. 
	NOTE: Do not over-dry the pellet as it will become difficult to resuspend.</li>
	<li>Optionally, assess the concentration of RNA by measuring absorbance at 260 nm.</li>
	<li>Optionally, assess the quality of the purified RNA by agarose gel electrophoresis20.</li>
	<li>Store samples at -80 &#176;C. 
	NOTE: The protocol can be paused here.</li>
</ol>
6. Preparation of chemically deacylated control sample
NOTE: To distinguish the band of aminoacylated tRNA from its deacylated counterpart on the Northern blot, a chemically deacylated aliquot is prepared by alkaline treatment. For most purposes, it is sufficient to deacylate a single sample for each Northern blot.
<ol>
	<li>Thaw the RNA sample on ice. Transfer a 4 &#181;L aliquot of the RNA sample to a new tube. Add 46 &#181;L Tris-HCl pH 9.0. Incubate for 2 h at 37 &#176;C.</li>
	<li>Add 15 &#181;L of 0.3 M sodium acetate at pH 4.5 and subsequently add 125 &#181;L of 96% ethanol. Precipitate RNA at -20 &#176;C for 1 h.</li>
	<li>Centrifuge for 30 min at 20,000 x g at 4 &#176;C. Completely remove supernatant and resuspend in 4 &#181;L cold 10 mM sodium acetate pH 4.5 and 1 mM EDTA. 
	NOTE: The protocol can be paused here, if the sample is stored at -80 &#176;C.</li>
</ol>
7. Gel electrophoresis and Northern blotting
<ol>
	<li>Mix 4 &#181;L sample (untreated or chemically deacylated) with 6 &#181;L loading buffer (0.1 M sodium acetate (pH 5.0), 8 M urea, 0.05% bromophenol blue, and 0.05% xylene cyanol). 
	NOTE: Remember to include the sample containing only spike-in cells, the chemically deacylated sample (and the sample without spike-in cells, if one was prepared).</li>
	<li>Load the samples onto a 0.4 mm thick 6.5% polyacrylamide gel (19:1 acrylam- ide:bisacrylamide) containing 8 M urea in 0.1 M sodium acetate buffer (pH 5.0). Use 60 cm long gels for proper separation of tRNA species.</li>
	<li>Separate the RNA by electrophoresis at 10 V/cm gel at 4 &#176;C, until the bromophenol blue reaches the bottom of the gel (~20 h).</li>
	<li>The area from the xylene cyanol dye and 20 cm down towards the bromophenol blue dye is used for blotting (the distance between the two dyes should be 25-30 cm). Carefully separate the two glass plates, so that the gel remains on one of them. A thin piece of filter paper is cut to the size of the desired blotting area and placed on top of the part of the gel that will be used for blotting. The parts of the gel that are not covered by filter paper are cut off and discarded.&#160; Use the filter paper to carefully lift the gel piece away from the glass plate. 
	NOTE: The large gel is very fragile so handle with care.</li>
	<li>A positively charged nylon membrane (e.g. Hybond N+ membrane) is placed on top of the gel and it is electroblotted at 20 V for 90 min using 40 mM Tris-acetate (pH 8.1), 2 mM EDTA as transfer buffer.</li>
	<li>Crosslink the RNA to the membrane using UV-light (0.12 J/cm2). 
	NOTE: The membrane can now be stored at room temperature and the protocol can be paused here.</li>
</ol>
8. Design and verification of probes
NOTE: Careful design and verification of Northern blot probes is crucial to obtain reliable results and avoid unwanted cross-hybridization with other RNA species.
<ol>
	<li>Use short synthetic oligonucleotides as probes. Design the probe sequence in a way that it is complementary to the most unique part of the sequence of the tRNA of interest - this is often the anticodon loop. 
	NOTE: A list of predicted tRNA sequences from various organisms can be found in the Genomic tRNA Database (GtRNAdb)21.</li>
	<li>Adjust the length of the sequence to obtain a predicted melting temperature of 55-65 &#176;C.</li>
	<li>BLAST (Basic Local Alignment Search Tool) the chosen sequence against the genome sequence of the bacterial strain used in the experiment, to verify that the sequence is unique to the selected tRNA, as described in22. 
	NOTE: Table 1 lists suggested probe sequences for several different tRNAs of E. coli K12. If the probe is specific for the tRNA of interest, only two bands are visible on the Northern blot (charged and uncharged tRNA) and only one band is visible in the lane with the deacylated sample (uncharged tRNA). If more than two bands are present or if further validation of probe specificity is desired, see Discussion.</li>
</ol>
9. Hybridization of Northern blot
NOTE: In the following steps, the membrane is sequentially hybridized with probes complementary to the desired specific tRNAs, including a probe specific for the reference tRNAselC.
<ol>
	<li>Place the membrane in a hybridization tube and add 6 mL hybridization solution (0.9 M NaCl, 0.05 M NaH2PO4 (pH 7.7), 5 mM EDTA, 0.5% (w/v) SDS, 100 mg/mL of sheared and denatured herring sperm DNA and 5x Denhardt&#39;s solution (100x Denhardt&#39;s solution = 2% bovine serum albumin, 2% polyvinylpyrrolidone 40, and 2% Ficoll)).</li>
	<li>Pre-hybridize the membrane in the buffer for 1 h rotating at 42 &#176;C. Add 30 pmol radioactive oligo-DNA probe 5&#39;-end-labeled with 32P (prepared as described by Sambrook et al.23) 
	NOTE: CAUTION, the high-energy beta emissions from 32P can present a substantial skin and eye dose hazard. Proper precaution should be taken when using 32P. Radioactive waste should be disposed of according to the appropriate institutional safety protocols.</li>
	<li>Incubate the probe with the membrane overnight rotating at 42 &#176;C. Remove the probe using a disposable 10 mL pipette. 
	NOTE: If stored in a freezer the probe can be reused.</li>
	<li>In the hybridization tube, wash the membrane shortly in 50 mL of 0.3 M NaCl, 30 mM sodium citrate, 0.1% (w/v) SDS at room temperature. 
	NOTE: This first wash contains much unbound probe and must be handled as highly radioactive.</li>
	<li>Repeat step 9.4 once, then move the membrane to a flat plastic container or tray including a lid. Cover the membrane with the washing solution (0.3 M NaCl, 30 mM sodium citrate, 0.1% (w/v) SDS) and incubate it for 30 min at room temperature.</li>
	<li>Change the washing solution and continue the washing until a satisfying signal to noise ratio has been obtained (often 3 - 4 changes of washing solution). Use a Geiger-M&#252;ller tube to detect how the noise to signal ratio changes by measuring the radiation from an empty area of the membrane and compare it to the area where the probe has hybridized specifically to tRNA.</li>
	<li>In order to be able to strip the probe from the membrane after exposure, ensure that (important) the membrane does not dry out before the stripping procedure. Place the membrane in a thin plastic bag or wrap it tightly using plastic wrap and seal it to be air-tight by tape or welding. Place the sealed membrane on a phosphorimaging screen. 
	NOTE: The required exposure time on the phosphorimaging screen is determined by the specific activity of the probe, the amount of RNA probed for, and the hybridization efficiency of the probe, and can vary greatly; from a few minutes to several days. With experience, the intensity of the radiation detected on the membrane with the Geiger-M&#252;ller tube gives a good indication of the required exposure time. For reliable quantification, it is important that the screen is not overexposed.</li>
	<li>Scan the phosphorimaging screen using a laser scanner and save the file in &#34;.gel&#34; format. 
	NOTE: A high signal to noise ratio is necessary for reliable quantification. Probing for a tRNA should result in two bands; one containing aminoacylated tRNA (slower migration) and one containing deacylated tRNA (faster migration). See Figure 2.</li>
</ol>
10. Stripping the membrane
NOTE: Before the membrane can be probed for another tRNA, the previous probe must first be removed by stripping the membrane.
<ol>
	<li>Heat enough stripping buffer (15 mM NaCl, 1.5 mM sodium citrate, 0.1% (w/v) SDS) to cover the membrane and pour it over the membrane. Incubate it for 15 min at room temperature.</li>
	<li>Repeat step 10.1 until no radiation can be detected with a Geiger-M&#252;ller tube; the membrane can now be re-hybridized following the steps described in Section 9.</li>
</ol>
11. Quantifying tRNA and charging levels
<ol>
	<li>Load the .gel file into the imaging software (see the Table of Materials). Draw a vertical line along each of the sample lanes in such a way that it spans most of the width of the lane but excludes the edges of the bands where the signal can be uneven, as shown in Figure 3A.</li>
	<li>Visualize counts from each lane by clicking &#34;Analysis&#34; -&#62; &#34;Create Graph&#34;. Manually define the two peaks representing the charged and the uncharged tRNA from each lane, as shown in Figure 3B.</li>
	<li>Obtain counts from each peak by clicking &#34;Analysis&#34; -&#62; &#34;Area Report&#34;, and record the area under each of the two peaks.</li>
	<li>When the blot has been probed for tRNAselC and at least one tRNA of interest, normalize the levels of the tRNA of interest to tRNAselC.
	<ol>
		<li>Calculate the proportion of tRNA originating from the spike-in cells in each lane and subtract from the total signal in that lane using the equation: 
		<img alt="Equation 3" src="/files/ftp_upload/56212/56212eq3.jpg" /> 
		NOTE: Here, SampletRNA = the signal obtained from a single lane of a membrane probed for the desired tRNA; SampleselC = the signal obtained from the same lane of the same membrane probed for tRNAselC; ReftRNA = the signal obtained from the lane containing only spike-in cell RNA of the same membrane probed for the desired tRNA; RefselC = the signal obtained from the lane containing only spike-in cell RNA of the same membrane probed for tRNAselC.</li>
		<li>To normalize, divide the corrected tRNA value from each lane by the tRNAselC signal from the same lane. 
		NOTE: The contribution to the tRNAselC signal from the sampled cells is negligible (See Figure 2, lane labeled &#34;-selC&#34;).</li>
	</ol>
	</li>
	<li>Calculate the tRNA charging level for each lane by dividing the signal of the peak representing charged tRNA with the sum of the signal from both peaks (charged + uncharged).</li>
</ol>

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The authors have nothing to disclose.

This protocol describes how to simultaneously measure the charging level of specific E. coli tRNAs and compare the relative levels of the tRNAs in different samples. The critical points of the protocol are 1) to handle the samples in such a way that the cellular tRNA charging levels are retained, 2) to normalize tRNA quantities in such a way that relative tRNA levels in different samples can be reliably compared, and 3) to ensure the specificity of the selected probes for the tRNAs of interest. These points are ...

In the following, we present a method for quantifying specific tRNAs and measuring their charging levels by Northern blotting. The method is based on a technique first developed by Varshney et al.1. By harvesting cells into trichloroacetic acid (TCA) and keeping the samples at 0 &#176;C throughout the RNA purification, the ester bond between the tRNA and the amino acid is conserved2,3,4. Aminoacylated tRNAs can be distinguished from their nonacylated counterparts by gel electrophoresis and Northern blotting, due to a decreased mobility of the aminoacylated tRNA in the gel, caused by the covalently bound amino acid5. Additionally, we present a protocol for normalizing tRNA quantities by addition of spike-in cells overexpressing the rarely used tRNAselC 4.
tRNAs are some of the most abundant molecules in the bacterial cell and an absolutely vital part of the translation machinery. tRNAs bind amino acids and transfer them to the translating ribosomes. The binding of amino acids to tRNAs (aminoacylation or charging) is facilitated by aminoacyl tRNA synthetases. The relative abundance of different charged tRNAs is important for ensuring the fidelity of protein synthesis, because underrepresentation of the cognate charged tRNA for a given messenger RNA (mRNA) codon increases the likelihood that a near-cognate tRNA will erroneously deliver its amino acid to the growing polypeptide chain on the ribosome6. The importance of charged tRNA is reflected in the extensive response of the E. coli cell to a severe drop in the charging levels of a tRNA; the stringent response. During the stringent response the synthesis of tRNAs, ribosomal RNAs and most mRNAs is lowered in favor of transcription of specific mRNAs associated with amino acid biosynthesis and stress survival, and the growth rate of the cells is lowered dramatically7. Furthermore, recent work by us and others has shown that E. coli actively degrades the majority of its tRNA in response to stresses that limit translation4,8, suggesting that adjustment of the tRNA levels may be important for coping with such stresses. Thus, reliable measurements of tRNA abundances and charging levels will be an important tool for fully understanding bacterial stress responses.
E.coli is commonly used to express recombinant protein and due to the differences in codon usage between species, suboptimal expression is a problem often faced9. This can be circumvented by expression of additional tRNAs needed to translate the recombinant mRNA10. Measurements of tRNA charging levels in such strains could guide troubleshooting efforts and help optimize protein expression.
This method also enables the detection of &#34;mischarging&#34;; a tRNA molecule aminoacylated with a non-cognate amino acid. Aminoacylation by different amino acids may cause a tRNA to migrate with slightly different velocities through polyacrylamide gels1,11. In some cases, the method can also be used to distinguish different modification patterns on otherwise identical tRNAs3.
Another established biochemical procedure for the investigation of tRNA charging levels is periodate oxidation. The method relies on the observation that aminoacylated tRNA is protected from periodate oxidation and uncharged tRNA is not. After periodate oxidation treatment the recharging of the tRNA is used to estimate the charging levels of the harvested RNA. However, the recharging of several tRNAs has been shown to be affected by the treatment thus providing some inaccuracy12. The method presented here measures charging directly from purified RNA, thus excluding any biases from chemical or enzymatic reactions. One limitation of this method is that only one tRNA species is detected at a time, so although the same Northern blot can be stripped and reprobed for multiple tRNAs, it is time-consuming and somewhat laborious to collect data on many tRNAs.
A reliable way of normalizing samples to each other is vital in studies where the goal is to compare the relative levels of a molecule across different samples. Here we introduce a normalization procedure for RNA where a small aliquot of E. coli cells overexpressing the rare tRNAselC is added as a spike-in to all the experimental samples prior to RNA purification. This method is useful not only when examining tRNAs but for relative quantification of any species of RNA when the experimental setup is such that no endogenous RNA can be trusted to be present at the same cellular concentration in all the samples. For example, the level of a &#34;housekeeping&#34; RNA-like ribosomal RNA is often used as an endogenous reference to compare the relative quantities of another RNA between different samples13. But this is of little use if the cellular concentration of the reference RNA varies between samples, as can be the case for ribosomal RNA if sampling occurs during a stress response15,16,17 or during entry into stationary phase17. The addition of spike-in cells to the experimental samples prior to RNA purification provides a solid and accurate way of normalization independent of the sampling setup. Another way of standardization is the addition of one or more spike-in RNAs to the experimental samples after RNA purification. However, this method does not account for any differences in RNA recovery between samples.
Using E. coli cells that overexpress the reference RNA as spike-in cells (see Figure 1) in an experiment on E. coli has the drawback that it results in addition of a small amount of exogenous E. coli total RNA to the samples. We correct for this addition by analyzing a sample containing only spike-in cells in parallel with the experimental samples (see the Northern blot in Figure 2, lane labeled &#34;selC&#34;). The protocol presented is developed for E. coli K-12 but is likely to be applicable for most bacterial species.

<table>
	<tbody>
		<tr>
			<td>Urea</td>
			<td>Merck</td>
			<td>57-13-6</td>
			<td>Purity &#62;99%</td>
		</tr>
		<tr>
			<td>Polyacrylamide</td>
			<td>Serva</td>
			<td>79-06-7</td>
			<td></td>
		</tr>
		<tr>
			<td>Bis (N,N&#39;-Methylene-bis-acrylamide)</td>
			<td>BIO-RAD</td>
			<td>161-0201</td>
			<td></td>
		</tr>
		<tr>
			<td>Trichloroacetic acid (TCA)</td>
			<td>Sigma</td>
			<td>76-03-9</td>
			<td></td>
		</tr>
		<tr>
			<td>Phenol</td>
			<td>Merck</td>
			<td>108-95-2</td>
			<td></td>
		</tr>
		<tr>
			<td>IPTG</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Acetate</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>EDTA</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Tris</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrochloric acid</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Chloride</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Citrate</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>SDS</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Herring Sperm DNA</td>
			<td>Sigma</td>
			<td>100403-24-5</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Polivinylpyrrolidone</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ficoll</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>P32 &#947;ATP</td>
			<td>Perkin Elmer</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>DNA oligos (probes)</td>
			<td>TAG Copenhagen</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Hybond N+ membrane</td>
			<td>GE Healthcare</td>
			<td>RPN203B</td>
			<td></td>
		</tr>
		<tr>
			<td>Crosslinker</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Electroblotter</td>
			<td>BIO-RAD</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Typhoon FLA 7000 Scanner</td>
			<td>GE Healthcare</td>
			<td>28955809</td>
			<td></td>
		</tr>
		<tr>
			<td>Spectrophotometer</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Hydridization oven</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Geiger-M&#252;ller tube</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphor imager screen</td>
			<td>GE Healthcare</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Hybridization tube</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Culture Flasks</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1.5 ml microcentrifuge tubes</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>20 ml centrifuge tubes</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Software</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ImageQuant</td>
			<td>GE Healthcare</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Excel</td>
			<td>Microsoft</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

quantification of the abundance and charging levels of transfer rnas in escherichia coli

Transfer RNA (tRNA) is an essential part of the translational machinery in any organism. tRNAs bind and transfer amino acids to the translating ribosome. The relative levels of different tRNAs, and the ratio of aminoacylated tRNA to total tRNA, known as the charging level, are important factors in determining the accuracy and speed of translation. Therefore, the abundance and charging levels of tRNAs are important variables to measure when studying protein synthesis, for example under various stress conditions. Here, we describe a method for harvesting tRNA and directly measuring both the relative abundance and the absolute charging level of specific tRNA species in Escherichia coli. The tRNA is harvested in such a way that the labile bond between the tRNA and its amino acid is preserved. The RNA is then subjected to gel electrophoresis and Northern blotting, which results in separation of the charged and uncharged tRNAs. The levels of specific tRNAs in different samples can be compared due to the addition of spike-in cells for normalization. Prior to RNA purification, we add 5% of E. coli cells that overproduce the rare tRNAselC to each sample. The amount of the tRNA species of interest in a sample is then normalized to the amount of tRNAselC in the same sample. Addition of spike-in cells prior to RNA purification has the advantage over addition of purified spike-in RNAs that it also accounts for any differences in cell lysis efficiency between samples.

Using the procedure described here, the abundance and charging levels of three tRNAs were measured in E. coli K-12 before and during amino acid starvation.
The arginine auxotroph strain NF9154 (thr leu his argH thi mtl supE44) was grown for at least ten generations in MOPS minimal medium supplemented with 0.4% glycerol, 50 &#181;g/mL threonine, leucine, arginine, and 5 &#181;g/mL histidi...

Watch this Scientific Journal Video about Quantification of the Abundance and Charging Levels of Transfer RNAs in Escherichia coli at JoVE.com

Quantification of the Abundance and Charging Levels of Transfer RNAs in Escherichia coli

Here we present a method for directly measuring transfer RNA charging levels from purified Escherichia coli RNA as well as a way to compare relative levels of transfer RNA, or any other short RNA, across different samples based on the addition of spike-in cells expressing a reference gene.

Quantification of the Abundance and Charging Levels of Transfer RNAs in Escherichia coli

Transfer RNA (tRNA) is an essential part of the translational machinery in any organism. tRNAs bind and transfer amino acids to the translating ribosome. ...

quantification-abundance-charging-levels-transfer-rnas-escherichia

Research

JoVE Journal

Biochemistry

9.2K Views.  University of Copenhagen. Here we present a method for directly measuring transfer RNA charging levels from purified Escherichia coli RNA as well as a way to compare relative levels of transfer RNA, or any other short RNA, across different samples based on the addition of spike-in cells expressing a reference gene.

Video: Quantification of the Abundance and Charging Levels of Transfer RNAs in Escherichia coli

Directed Protein Packaging within Outer Membrane Vesicles from Escherichia coli: Design, Production and Purification

An increasing interest in applying synthetic biology techniques to program outer membrane vesicles (OMV) are leading to some very interesting and unique applications for OMV where traditional nanoparticles are proving too difficult to synthesize. To date, all Gram-negative bacteria have been shown to produce OMV demonstrating packaging of a variety of cargo that includes small molecules, peptides, proteins and genetic material. Based on their diverse cargo, OMV are implicated in many biological processes ranging from cell-cell communication to gene transfer and delivery of virulence factors depending upon which bacteria are producing the OMV. Only recently have bacterial OMV become accessible for use across a wide range of applications through the development of techniques to control and direct packaging of recombinant proteins into OMV. This protocol describes a method for the production, purification, and use of enzyme packaged OMV providing for improved overall production of recombinant enzyme, increased vesiculation, and enhanced enzyme stability. Successful utilization of this protocol will result in the creation of a bacterial strain that simultaneously produces a recombinant protein and directs it for OMV encapsulation through creating a synthetic linkage between the recombinant protein and an outer membrane anchor protein. This protocol also details methods for isolating OMV from bacterial cultures as well as proper handling techniques and things to consider when adapting this protocol for use for other unique applications such as: pharmaceutical drug delivery, medical diagnostics, and environmental remediation.

Directed Protein Packaging within Outer Membrane Vesicles from Escherichia coli: Design, Production and Purification

A protocol for the production, purification, and use of enzyme packaged outer membrane vesicles (OMV) providing for enhanced enzyme stability for implementation across diverse applications is presented.

An increasing interest in applying synthetic biology techniques to program outer membrane vesicles (OMV) are leading to some very interesting and unique ...

Detection of the pH-dependent Activity of Escherichia coli Chaperone HdeB In Vitro and In Vivo

Bacteria are frequently exposed to environmental changes, such as alterations in pH, temperature, redox status, light exposure or mechanical force. Many of these conditions cause protein unfolding in the cell and have detrimental impact on the survival of the organism. A group of unrelated, stress-specific molecular chaperones have been shown to play essential roles in the survival of these stress conditions. While fully folded and chaperone-inactive before stress, these proteins rapidly unfold and become chaperone-active under specific stress conditions. Once activated, these conditionally disordered chaperones bind to a large number of different aggregation-prone proteins, prevent their aggregation and either directly or indirectly facilitate protein refolding upon return to non-stress conditions. The primary approach for gaining a more detailed understanding about the mechanism of their activation and client recognition involves the purification and subsequent characterization of these proteins using in vitro chaperone assays. Follow-up in vivo stress assays are absolutely essential to independently confirm the obtained in vitro results.
This protocol describes in vitro and in vivo methods to characterize the chaperone activity of E. coli HdeB, an acid-activated chaperone. Light scattering measurements were used as a convenient read-out for HdeB&#39;s capacity to prevent acid-induced aggregation of an established model client protein, MDH, in vitro. Analytical ultracentrifugation experiments were applied to reveal complex formation between HdeB and its client protein LDH, to shed light into the fate of client proteins upon their return to non-stress conditions. Enzymatic activity assays of the client proteins were conducted to monitor the effects of HdeB on pH-induced client inactivation and reactivation. Finally, survival studies were used to monitor the influence of HdeB&#39;s chaperone function in vivo.

Detection of the pH-dependent Activity of Escherichia coli Chaperone HdeB In Vitro and In Vivo

This study describes biophysical, biochemical and molecular techniques to characterize the chaperone activity of Escherichia coli HdeB under acidic pH conditions. These methods have been successfully applied for other acid-protective chaperones such as HdeA and can be modified to work for other chaperones and stress conditions.

Bacteria are frequently exposed to environmental changes, such as alterations in pH, temperature, redox status, light exposure or mechanical force. Many ...

Quantification of Hypopigmentation Activity In Vitro

This study presents laboratory methods for the quantification of hypopigmentation activity in vitro. Melanin, the major pigment in melanocytes, is synthesized in response to multiple cellular and environmental factors. Melanin protects skin cells from ultraviolet damage, but also has biophysical and biochemical functions. Excessive production or accumulation of melanin in melanocytes can cause dermatological problems, such as freckles, dark spots, melasma, and moles. Therefore, the control of melanogenesis with hypopigmentation agents is important in individuals with clinical or cosmetic needs. Melanin is primarily synthesized in the melanosomes of melanocytes in a complex biochemical process called melanogenesis, which is influenced by extrinsic and intrinsic factors, such as hormones, inflammation, age, and ultraviolet light exposure. We describe three methods to determine the hypopigmentation activity of chemicals or natural substances in melanocytes: measurement of the 1) cellular tyrosinase activity and 2) melanin content, and 3) staining and quantifying cellular melanin with image analysis.
In melanogenesis, tyrosinase catalyzes the rate-limiting step that converts L-tyrosine into 3,4-dihydroxyphenylalanine (L-DOPA) and then into dopaquinone. Therefore, the inhibition of tyrosinase is a primary hypopigmentation mechanism. In cultured melanocytes, tyrosinase activity can be quantified by adding L-DOPA as a substrate and measuring dopaquinone production by spectrophotometry. Melanogenesis can also be measured by quantifying the melanin content. The melanin-containing cellular fraction is extracted with NaOH and melanin is quantified spectrophotometrically. Finally, the melanin content can be quantified by image analysis following Fontana-Masson staining of melanin. Although the results of these in vitro assays may not always be reproduced in human skin, these methods are widely used in melanogenesis research, especially as the initial step to identify potential hypopigmentation activity. These methods can also be used to assess melanocyte activity, growth, and differentiation. Consistent results with the three different methods ensure the validity of the effects.

We describe three experimental methods for evaluating the hypopigmentation activity of chemicals in vitro: quantification of 1) cellular tyrosinase activity and 2) melanin content, and 3) measurement of melanin by cellular melanin staining and image analysis.

This study presents laboratory methods for the quantification of hypopigmentation activity in vitro. Melanin, the major pigment in melanocytes, is ...

Large-scale Production of Recombinant RNAs on a Circular Scaffold Using a Viroid-derived System in Escherichia coli

With increasing interest in RNA biology and the use of RNA molecules in sophisticated biotechnological applications, the methods to produce large amounts of recombinant RNAs are limited. Here, we describe a protocol to produce large amounts of recombinant RNA in Escherichia coli based on co-expression of a chimeric molecule that contains the RNA of interest in a viroid scaffold and a plant tRNA ligase. Viroids are relatively small, non-coding, highly base-paired circular RNAs that are infectious to higher plants. The host plant tRNA ligase is an enzyme recruited by viroids that belong to the family Avsunviroidae, such as Eggplant latent viroid (ELVd), to mediate RNA circularization during viroid replication. Although ELVd does not replicate in E. coli, an ELVd precursor is efficiently transcribed by the E. coli RNA polymerase and processed by the embedded hammerhead ribozymes in bacterial cells, and the resulting monomers are circularized by the co-expressed tRNA ligase reaching a remarkable concentration. The insertion of an RNA of interest into the ELVd scaffold enables the production of tens of milligrams of the recombinant RNA per liter of bacterial culture in regular laboratory conditions. A main fraction of the RNA product is circular, a feature that facilitates the purification of the recombinant RNA to virtual homogeneity. In this protocol, a complementary DNA (cDNA) corresponding to the RNA of interest is inserted in a particular position of the ELVd cDNA in an expression plasmid that is used, along the plasmid to co-express eggplant tRNA ligase, to transform E. coli. Co-expression of both molecules under the control of strong constitutive promoters leads to production of large amounts of the recombinant RNA. The recombinant RNA can be extracted from the bacterial cells and separated from the bulk of bacterial RNAs taking advantage of its circularity.

Large-scale Production of Recombinant RNAs on a Circular Scaffold Using a Viroid-derived System in Escherichia coli

Here, we present a protocol to produce large amounts of recombinant RNA in Escherichia coli by co-expressing a chimeric RNA that contains the RNA of interest in a viroid scaffold and a plant tRNA ligase. The main product is a circular molecule that facilitates purification to homogeneity.

With increasing interest in RNA biology and the use of RNA molecules in sophisticated biotechnological applications, the methods to produce large amounts ...

Site-Directed Mutagenesis for In Vitro and In Vivo Experiments Exemplified with RNA Interactions in Escherichia Coli

Site-directed mutagenesis is a technique used to introduce specific mutations in DNA to investigate the interaction between small non-coding ribonucleic acid (sRNA) molecules and target messenger RNAs (mRNAs). In addition, site-directed mutagenesis is used to map specific protein binding sites to RNA. A 2-step and 3-step PCR based introduction of mutations is described. The approach is relevant to all protein-RNA and RNA-RNA interaction studies. In short, the technique relies on designing primers with the desired mutation(s), and through 2 or 3 steps of PCR synthesizing a PCR product with the mutation. The PCR product is then used for cloning. Here, we describe how to perform site-directed mutagenesis with both the 2- and 3-step approach to introduce mutations to the sRNA, McaS, and the mRNA, csgD, to investigate RNA-RNA and RNA-protein interactions. We apply this technique to investigate RNA interactions; however, the technique is applicable to all mutagenesis studies (e.g., DNA-protein interactions, amino-acid substitution/deletion/addition). It is possible to introduce any kind of mutation except for non-natural bases but the technique is only applicable if a PCR product can be used for downstream application (e.g., cloning and template for further PCR).

Site-Directed Mutagenesis for In Vitro and In Vivo Experiments Exemplified with RNA Interactions in Escherichia Coli

Site-directed mutagenesis is a technique used to introduce specific mutations in deoxyribonucleic acid (DNA). This protocol describes how to do site-directed mutagenesis with a 2-step and 3-step polymerase chain reaction (PCR) based approach, which is applicable to any DNA fragment of interest.

Site-directed mutagenesis is a technique used to introduce specific mutations in DNA to investigate the interaction between small non-coding ribonucleic ...

Use of Alu Element Containing Minigenes to Analyze Circular RNAs

In addition to linear mRNAs, many eukaryotic genes generate circular RNAs. Most circular RNAs are generated by joining a 5&#39; splice site with an upstream 3&#39; splice site within a pre-mRNA, a process called back-splicing. This circularization is likely aided by secondary structures in the pre-mRNA that bring the splice sites into close proximity. In human genes, Alu elements are thought to promote these secondary RNA structures, as Alu elements are abundant and exhibit base complementarities with each other when present in opposite directions in the pre-mRNA. Here, we describe the generation and analysis of large, Alu element containing reporter genes that form circular RNAs. Through optimization of cloning protocols, reporter genes with up to 20 kb insert length can be generated. Their analysis in co-transfection experiments allows the identification of regulatory factors. Thus, this method can identify RNA sequences and cellular components involved in circular RNA formation.

Use of Alu Element Containing Minigenes to Analyze Circular RNAs

We clone and analyze reporter genes generating circular RNAs. These reporter genes are larger than constructs to analyze linear splicing and contain Alu elements. To investigate the circular RNAs, the constructs are transfected into cells and resulting RNA is analyzed using RT-PCR after removal of linear RNA.

In addition to linear mRNAs, many eukaryotic genes generate circular RNAs. Most circular RNAs are generated by joining a 5&#39; splice site with an ...

Synthesis of a Deuterated Standard for the Quantification of 2-Arachidonoylglycerol in Caenorhabditis elegans

This work presents a method to prepare an analytical standard to analyze 2-arachidonoyl glycerol (2-AG) qualitatively and quantitatively by liquid chromatography-electrospray Ionization-tandem mass spectrometry (LC-ESI-MS/MS). Endocannabinoids are conserved lipid mediators that regulate multiple biological processes in a variety of organisms. In C. elegans, 2-AG has been found to possess different roles, including modulation of dauer formation and cholesterol metabolism. This report describes a method to overcome the difficulties associated with the costs and stability of deuterated standards required for 2-AG quantification. The procedure for the synthesis of the standard is simple and can be performed in any laboratory, without the need for organic synthesis expertise or special equipment. In addition, a modification of Folch&#39;s method to extract the deuterated standard from C. elegans culture is described. Finally, a quantitative and analytic method to detect 2-AG using the stable isotopically labeled analog 1-AG-d5 is described, which provides reliable results in a fast-chromatographic run. The procedure is useful for studying the multiple roles of 2-AG in C. elegans while also being applicable to other studies of metabolites in different organisms.

Synthesis of a Deuterated Standard for the Quantification of 2-Arachidonoylglycerol in Caenorhabditis elegans

This work describes a robust and straightforward method to detect and quantify the endocannabinoid 2-arachidonoylglycerol (2-AG) in C. elegans. An analytical deuterated standard us prepared and used for the quantification of 2-AG by isotopic dilution and liquid chromatography-electrospray ionization-tandem mass spectrometry (LC-ESI-MS/MS).

This work presents a method to prepare an analytical standard to analyze 2-arachidonoyl glycerol (2-AG) qualitatively and quantitatively by liquid ...

Characterization of Membrane Transporters by Heterologous Expression in E. coli and Production of Membrane Vesicles

Several methods have been developed to functionally characterize novel membrane transporters. Polyamines are ubiquitous in all organisms, but polyamine exchangers in plants have not been identified. Here, we outline a method to characterize polyamine antiporters using membrane vesicles generated from the lysis of Escherichia coli cells heterologously expressing a plant antiporter. First, we heterologously expressed AtBAT1 in an E. coli strain deficient in polyamine and arginine exchange transporters. Vesicles were produced using a French press, purified by ultracentrifugation and utilized in a membrane filtration assay of labeled substrates to demonstrate the substrate specificity of the transporter. These assays demonstrated that AtBAT1 is a proton-mediated transporter of arginine, &#947;-aminobutyric acid (GABA), putrescine and spermidine. The mutant strain that was developed for the assay of AtBAT1 may be useful for the functional analysis of other families of plant and animal polyamine exchangers. We also hypothesize that this approach can be used to characterize many other types of antiporters, as long as these proteins can be expressed in the bacterial cell membrane. E. coli is a good system for the characterization of novel transporters, since there are multiple methods that can be employed to mutagenize native transporters.

Characterization of Membrane Transporters by Heterologous Expression in E. coli and Production of Membrane Vesicles

We describe a method for the characterization of proton-driven membrane transporters in membrane vesicle preparations produced by heterologous expression in E. coli and lysis of cells using a French press.

Several methods have been developed to functionally characterize novel membrane transporters. Polyamines are ubiquitous in all organisms, but polyamine ...

Quantification of Protein Interaction Network Dynamics using Multiplexed Co-Immunoprecipitation

Dynamic protein-protein interactions control cellular behavior, from motility to DNA replication to signal transduction. However, monitoring dynamic interactions among multiple proteins in a protein interaction network is technically difficult. Here, we present a protocol for Quantitative Multiplex Immunoprecipitation (QMI), which allows quantitative assessment of fold changes in protein interactions based on relative fluorescence measurements of Proteins in Shared Complexes detected by Exposed Surface epitopes (PiSCES). In QMI, protein complexes from cell lysates are immunoprecipitated onto microspheres, and then probed with a labeled antibody for a different protein in order to quantify the abundance of PiSCES. Immunoprecipitation antibodies are conjugated to different MagBead spectral regions, which allows a flow cytometer to differentiate multiple parallel immunoprecipitations and simultaneously quantify the amount of probe antibody associated with each. QMI does not require genetic tagging and can be performed using minimal biomaterial compared to other immunoprecipitation methods. QMI can be adapted for any defined group of interacting proteins, and has thus far been used to characterize signaling networks in T cells and neuronal glutamate synapses. Results have led to new hypothesis generation with potential diagnostic and therapeutic applications. This protocol includes instructions to perform QMI, from the initial antibody panel selection through to running assays and analyzing data. The initial assembly of a QMI assay involves screening antibodies to generate a panel, and empirically determining an appropriate lysis buffer. The subsequent reagent preparation includes covalently coupling immunoprecipitation antibodies to MagBeads, and biotinylating probe antibodies so they can be labeled by a streptavidin-conjugated fluorophore. To run the assay, lysate is mixed with MagBeads overnight, and then beads are divided and incubated with different probe antibodies, and then a fluorophore label, and read by flow cytometry. Two statistical tests are performed to identify PiSCES that differ significantly between experimental conditions, and results are visualized using heatmaps or node-edge diagrams.

Quantitative Multiplex Immunoprecipitation (QMI) uses flow cytometry for sensitive detection of differences in the abundance of targeted protein-protein interactions between two samples. QMI can be performed using a small amount of biomaterial, does not require genetically engineered tags, and can be adapted for any previously defined protein interaction network.

Dynamic protein-protein interactions control cellular behavior, from motility to DNA replication to signal transduction. However, monitoring dynamic ...

Quantification of Coenzyme A in Cells and Tissues

Emerging research has revealed that the cellular coenzyme A (CoA) supply can become limiting with a detrimental impact on growth, metabolism and survival. Measurement of cellular CoA is a challenge due to its relatively low abundance and the dynamic conversion of free CoA to CoA thioesters that, in turn, participate in numerous metabolic reactions. A method is described that navigates through potential pitfalls during sample preparation to yield an assay with a broad linear range of detection that is suitable for use in many biomedical laboratories.

This method describes sample preparation from cultured cells and animal tissues, extraction and derivatization of coenzyme A in the samples, followed by high pressure liquid chromatography for purification and quantification of the derivatized coenzyme A by absorbance or fluorescence detection.

Emerging research has revealed that the cellular coenzyme A (CoA) supply can become limiting with a detrimental impact on growth, metabolism and survival. ...