ALS would like to acknowledge the financial support of EMBO Installation Grants IG 3914, and POIR. 04.04.00-00-3E9C/17-00 carried out within the First TEAM programme of the Foundation for Polish Science co-financed by the European Union under the European Regional Development Fund.

1. Sample collection
<ol>
	<li>Prepare a bacterial culture. We recommend a culture volume of 100 mL per sample.</li>
	<li>Prepare equipment and reagents for sample collection: two scoopulas per sample, sterile 0.45 &#181;m mixed cellulose esters membrane (MCE) filters, 50 mL sterile tubes, liquid nitrogen, 50 mg/mL chloramphenicol in 70% (vol/vol) ethanol (optional). Puncture the lid of 50 mL tubes to allow liquid nitrogen evaporation and prevent the explosion of closed tubes. Decontaminate scoopulas with laboratory detergent and 70% ethanol.</li>
	<li>Prewarm the filtering equipment and one of the scoopulas to growth temperature of the bacterial culture. 
	NOTE: We recommend an all-glass filtration apparatus with a funnel, a fritted base, a ground joint flask and a 47 mm &#216; spring clamp.</li>
	<li>Before sample harvesting, add chloramphenicol into the bacterial culture to the final concentration of 100 &#181;g/mL and incubate 1 minute (optional).</li>
	<li>Pour liquid nitrogen into 50 mL sterile tube and place the second scoopula inside the tube to cool. 
	CAUTION! Liquid nitrogen can cause closed containers to explode due to the pressure change upon evaporation. To prevent this, it is necessary to create a few holes in the lid of 50 mL tubes to allow liquid nitrogen evaporation.</li>
	<li>Collect cells by filtration. Stop filtering when medium has passed through the membrane, but do not allow the filter to dry completely.</li>
	<li>Collect bacterial pellets by rapidly scraping the cells off of the filter disc using a prewarmed scoopula. Immediately place the entire scoopula with the harvested cells in the 50 mL tube filled with liquid nitrogen. The harvested pellet should be completely covered in liquid nitrogen.</li>
	<li>Let the pellet freeze thoroughly and dislodge frozen cells using a previously cooled second scoopula. Close the punctured lid and transfer the tube to -80&#176;C. STOP point 
	&#8203;NOTE: Disinfect scoopulas after each sample harvesting.</li>
</ol>
2. Cell lysis
<ol>
	<li>Prepare 0.1 M GMP-PNP in 10 mM Tris pH 8, DNase I and 1.5 mL reaction tubes for lysates and keep them on ice. Cool the centrifuge to 4 &#176;C.</li>
	<li>Prepare lysis buffer (20 mM TRIS pH 7.6, 10 mM MgAcet, 150 mM KAcet, 0.4% TRITON X-100, 6 mM &#946;-mercaptoethanol, 5 mM CaCl2 and optional 1 mM chloramphenicol). Aliquot 500 &#181;L of the lysis buffer per sample into 1.5 mL reaction tubes and place them on ice.</li>
	<li>Decontaminate mortar and pestle with a laboratory detergent and 70% ethanol, and dry them. Cool mortar and pestle by pouring liquid nitrogen into the mortar.</li>
	<li>Transfer frozen pellet into the pre-chilled mortar and grind it to a powder. With a spatula, add approximately 1 volume of aluminum oxide and continue grinding. Keep mortar, pestle and cells cool by pouring liquid nitrogen when needed, do not let the contents of the mortar to thaw.</li>
	<li>Just before using lysis buffer, add GMP-PNP and DNase I into the lysis buffer aliquot to the final concentration of 2 mM and 100 U/mL respectively. Transfer the solution into the mortar with cells and aluminum oxide and continue grinding. Let the lysate thaw slowly while grinding and transfer the mixture into the pre-cooled 1.5 mL reaction tube and immediately place on ice.</li>
	<li>Centrifuge lysates at 20 000 x g for 5 minutes at 4 &#176;C. Transfer supernatants into new, pre-cooled 1,5 mL reaction tubes and keep them on ice.</li>
	<li>Measure the RNA concentration in each sample with a NanoDrop. Use 1:10 dilutions of samples and 1:10 dilution of lysis buffer in nuclease-free water as a blank. 
	NOTE: Be aware that chloramphenicol shows significant absorption at 260 nm.</li>
	<li>Divide each lysate into two portions: one for RIBO-seq (0.5 - 1 mg of RNA) and the second for RNA-seq (the rest).</li>
	<li>Clean the samples for RNA-seq using a commercially available RNA clean-up kit according to the manufacturer&#39;s protocol. 
	&#8203;NOTE: We recommend using Zymo RNA Clean &#38; Concentrate -25 kit (see Table of Materials). We also recommend adding 4.5 volumes of ethanol (compared to sample volume) into the mix of sample and Binding Buffer for ribosomal footprints and fragmented mRNA, in order to increase the purification efficiency of short RNA fragments.</li>
	<li>Store the samples intended for RNA-seq at -80 &#176;C. The procedure for RNA-seq will continue from step 5.</li>
</ol>
3. MNase digestion of samples for RIBO-seq
<ol>
	<li>To 1 mg of RNA add 3.8 &#181;L of 187.5 U/&#181;L MNase in 10 mM Tris pH 8 and top it up with lysis buffer to the total volume of 500 &#181;L.</li>
	<li>Incubate at 25 &#176;C, 300 RPM, for 45 minutes in a thermomixer.</li>
	<li>Clean the samples with a commercially available RNA clean up kit (as in step 2.9).</li>
	<li>Store the samples for RIBO-seq at -80 &#176;C (STOP point) or proceed to size selection.</li>
</ol>
4. Polyacrylamide gel electrophoresis (PAGE) and size selection of samples for RIBO-seq
<ol>
	<li>Prepare a 15% polyacrylamide-TBE gel with 8 M urea and place it in a tank with TBE buffer. Pre-run for minimum 10 minutes at a constant voltage of 200 V.</li>
	<li>Mix samples with TBE-Urea Sample Buffer, denature at 95 &#176;C for 1 minute and place them immediately on ice.</li>
	<li>Wash out the urea by injecting TBE buffer into gel wells using a syringe. Load the samples leaving one well space between them to separate each sample and prevent cross-contamination. Use 29 nt oligonucleotide and a mix of 26 nt and 32 nt oligonucleotides as markers. Run electrophoresis at a constant voltage of 180 V.</li>
	<li>Prepare sterile buffer for overnight incubation (5 mM EDTA, 10 mM NaOAc pH 5).</li>
	<li>After electrophoresis, stain the gel in a SYBR Gold-bath for about 2-3 minutes and rinse the gel with nuclease-free water.</li>
	<li>Excise fragments of gel between 26 and 32 nt with the sterile needles or the&#160;razor blades and place the gel fragments in separate 1.5 mL tubes for each sample. Change the needle or the razor blade between the samples.</li>
	<li>Add 200 &#181;L of the overnight incubation buffer to each reaction tube.</li>
	<li>Incubate the samples at 10 &#176;C, 1000 RPM overnight in a thermomixer.</li>
	<li>The next day, clean the samples as in step 2.9. Elute the footprints in 80 &#181;L of nuclease-free water.</li>
	<li>Store the samples at -80 &#176;C. STOP point. The procedure for RIBO-seq will continue from step 7.</li>
</ol>
5. Purification of bacterial mRNA by rRNA depletion from samples for RNA-seq
<ol>
	<li>Remove rRNA from samples with a bacterial mRNA purification kit (e.g., Invitrogen MICROBExpress). Follow the manufacturer&#39;s protocol.</li>
	<li>Clean the samples as in step 2.9. Elute the mRNA in 50 &#181;L of nuclease-free water.</li>
	<li>Store the samples for RNA-seq at -80 &#176;C (STOP point) or continue to alkaline fragmentation.</li>
</ol>
6. Alkaline fragmentation of samples for RNA-seq
<ol>
	<li>Prepare alkaline hydrolysis buffer (mix 220 &#181;L of 0.1 M NaHCO3, 30 &#181;L of 0.1 M Na2CO3 and 1 &#181;L of 0.5 M EDTA). Mix 1 volume (50 &#181;L) of alkaline buffer with 1 volume (50 &#181;L) of the sample and incubate at 95 &#176;C for 25 minutes.</li>
	<li>Add 5 &#181;L of 3 M NaOAc pH 5.5 to stop the reaction.</li>
	<li>Clean the samples as in step 2.9 and elute the samples in 80 &#181;L of nuclease-free water.</li>
	<li>Possible STOP point: store RNA-seq samples at -80 &#176;C. However, we recommend to go directly to step 7 in order to avoid unnecessary freezing and thawing.</li>
</ol>
7. Dephosphorylation and phosphorylation of samples for both RNA- and RIBO-seq
<ol>
	<li>Add 10 &#181;L of 10x reaction buffer PNK and 5 &#181;L T4 PNK to each sample. Incubate at 37 &#176;C for 1.5 hour in order to dephosphorylate the 3&#39; ends.</li>
	<li>Add 3 &#181;L of 1 mM ATP and incubate at 37 &#176;C for 1 hour in order to phosphorylate the 5&#39; ends.</li>
	<li>Clean the samples as in step 2.9, elute in 30 &#181;L nuclease-free water.</li>
	<li>Store the samples at -80 &#176;C. STOP point.</li>
</ol>
8. Library preparation using NEBNext Multiplex Small RNA Library Prep Set for Illumina
<ol>
	<li>Prepare 100-1000 ng of input RNA (obtained mRNA fragments and ribosomal footprints) in 6 &#181;L of nuclease-free water and add 1 &#181;L of 3&#39; SR Adaptor. Incubate according to the manufacturer&#39;s protocol.</li>
	<li>Add 10 &#181;L of 3&#39; Ligation Reaction Buffer (2X) and 3 &#181;L of 3&#39; Ligation Enzyme Mix and incubate at 37 &#176;C for 2.5 hour, instead of 1 hour at 25 &#176;C as the manufacturer&#39;s protocol specifies.</li>
	<li>Hybridize the Reverse Transcription Primer according to the manufacturer&#39;s protocol with a modified program: 5 minutes at 75 &#176;C, 30 minutes at 37 &#176;C and hold at 4 &#176;C.</li>
	<li>Ligate the 5&#180; SR Adaptor. Follow the manufacturer&#39;s protocol with one change: perform the incubation at 37 &#176;C for 2.5 hour, instead of 1 hour at 25 &#176;C.</li>
	<li>Perform Reverse Transcription according to the manufacturer&#39;s protocol.</li>
	<li>Possible STOP point: incubate the samples containing synthesized cDNAs at 70 &#176;C for 15 minutes to inactivate the reverse transcriptase and store them at -80 &#176;C. However, we recommend to proceed directly to the next steps to avoid unnecessary freezing and thawing of the samples.</li>
	<li>Store half of each cDNA library at -80 &#176;C as a back-up.</li>
	<li>Perform PCR amplification according to the manufacturer&#39;s protocol. Use half of the reaction mix. Store the obtained libraries at -80 &#176;C. STOP point.</li>
</ol>
9. Size selection of cDNA libraries using PAGE
<ol>
	<li>Prepare 6% polyacrylamide-TBE gel and place it in a tank with TBE buffer. Pre-run for minimum 10 minutes at a constant voltage of 200 V.</li>
	<li>Mix the samples with Gel Loading Dye, Blue (6X) and load them on the gel. Use Quick-Load pBR322 DNA-MspI Digest as a ladder. At the beginning, run electrophoresis at a constant voltage of 120 V to allow DNA to enter the gel and then change the voltage to 180 V.</li>
	<li>When electrophoresis is finished, stain the gel in a SYBR Gold-bath for about 2-3 minutes and rinse the gel with nuclease-free water.</li>
	<li>Excise gel fragments containing the libraries. For RNA-seq samples excise between 135-180 nt and for RIBO-seq between 135-170 nt. Use sterile needle or razor blade and place the excised gel fragments in separate 1.5 mL reaction tubes. Remember to change the needle or razor blade between the samples. 
	NOTE: The adapters and barcode from NEBNext Multiplex Small RNA Library Prep Set for Illumina have 119 nt in total which determines the lower excision cut-off.</li>
	<li>Add 100 &#181;L of nuclease-free water to each excised gel fragment.</li>
	<li>Incubate the gel fragments at 10 &#176;C, 450 RPM overnight in a thermomixer.</li>
	<li>Clean and concentrate the cDNA libraries with a commercial DNA clean-up kit according to the manufacturer&#39;s protocol. 
	NOTE: We recommend using Zymo DNA Clean &#38; Concentrate -5 kit (see Table of Materials).</li>
	<li>Store purified cDNA libraries at -80 &#176;C. STOP point.</li>
</ol>

The authors have nothing to disclose.

The key technical challenge of the ribosome profiling is the need to rapidly inhibit translation in order to capture a snapshot of ribosomes on mRNAs at a particular physiological state of interest. To accomplish this, translation inhibitors, rapid harvesting and flash freezing in liquid nitrogen are commonly used. Applying antibiotics is optional since they can cause artifacts. Chloramphenicol is a commonly used drug to arrest elongating ribosomes in bacterial RIBO-seq. However, it does not prevent initiation, resulting...

The ribosome profiling technique (RIBO-seq) was developed in the laboratory of Jonathan Weissman at the University of California, San Francisco1. In comparison to other methods used to study gene expression at the translational level, RIBO-seq focuses on each ribosome binding to mRNA and provides information about its location and the relative number of ribosomes on a transcript. It enables monitoring the process of protein synthesis in vivo and can provide single codon resolution and accuracy allowing the measurement of the ribosome density on both, the individual mRNA and along the entire transcriptome in the cell. At the foundation of the RIBO-seq technique lies the fact that during translation the ribosome binds the mRNA molecule and thus protects the buried fragment of the transcript from a ribonuclease digestion. Upon addition of the ribonuclease, the unprotected mRNA is digested and the fragments enclosed by ribosomes - typically of ~28-30 nt long - remain intact. These fragments, called ribosomal footprints (RF), can then be isolated, sequenced and mapped onto the transcript they originated from resulting in the detection of the exact position of the ribosomes. In fact, the ribosome ability to protect mRNA fragments has been used since the 1960s to study ribosomal binding and translation initiation sites (TIS)2,3,4. However, with the advancement in deep sequencing technology, RIBO-seq has become a gold standard for translation monitoring5 which, through the ribosome engagement, can provide a genome-wide information on protein synthesis6. Ribosome profiling filled the technological gap that existed between quantifying the transcriptome and the proteome6.
To conduct ribosome profiling we need to obtain cell lysate of the organism that had grown under the investigated conditions. Disrupting these conditions during cell collection and lysis may provide unreliable data. To prevent this, translation inhibitors, rapid harvesting and flash freezing in liquid nitrogen are commonly used. Cells can be lysed by cryogenic grinding in a mechanical homogenizer like a mixer mill7,8 or a bead beater9, and by trituration through a pipette10 or with a needle11. The lysis buffer can be added just before or shortly after pulverization of the cells. In our protocol we use liquid nitrogen to precool mortar and pestle, as well as aluminum oxide as a gentler approach to disruption of the bacterial cell wall, which prevents RNA shearing often encountered when methods such as sonification are applied. After pulverization, we add an ice-cold lysis buffer into the cooled contents of the mortar. Selection of an appropriate lysis buffer is important for obtaining the best resolution of ribosomal footprints. Since ionic strength affects both the RF size and the reading frame precision, it is currently recommended to use lysis buffers with low ionic strength and buffer capacity, even if it appears that buffer composition does not affect ribosomal occupancy on mRNAs11,12. Important components of the lysis buffer are magnesium ions, the presence of which prevents dissociation of the ribosomal subunits and inhibits spontaneous conformational changes in the bacterial ribosomes11,13. Calcium ions also play a significant role and are essential for the activity of micrococcal nuclease (MNase) used in the bacterial ribosome profiling method14. Addition of guanosine 5&#8242;-[&#946;,&#947;-imido]triphosphate (GMP-PNP), a non-hydrolyzable analog of GTP, together with chloramphenicol inhibits translation during lysis15.
When the lysate is obtained, it is clarified by centrifugation and divided into two portions, each for a RIBO-seq and a high-throughput total mRNA sequencing (RNA-seq) since they are performed simultaneously (Figure 1). RNA-seq provides a point of reference which enables the comparison of data from both RIBO-seq and RNA-seq during data analysis. The investigated translatome is defined by normalization of ribosomal footprints to mRNA abundance16. Data from RNA-seq can also help identify cloning or sequencing artifacts17.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62544/62544fig01.jpg" /> 
Figure 1. Schematics of mRNA sample preparation for RIBO-seq and RNA-seq. For RIBO-seq library preparation, RNA is digested with MNase (A), followed by the size selection of RF of ~28-30 nt length (B); for RNA-seq RNA is isolated (C), depleted of rRNA (D), and the resulting mRNA is randomly fragmented into fragments of varying lengths (E). <a href="https://www.jove.com/files/ftp_upload/62544/62544fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Initial steps of the procedure of sample preparation for RIBO-seq and RNA-seq differ slightly (Figure 1). For the ribosomal profiling, the lysate needs to be digested by a specific endonuclease to degrade the mRNA molecules not protected by the ribosomes. In standard protocols, the obtained monosomes are recovered by a sucrose cushion ultracentrifugation or a sucrose gradient ultracentrifugation8,14. In this article, we show that this step is not necessary to isolate RF required for the RIBO-seq in bacteria, likewise for eukaryotic cells18, and that size selection of the appropriate length mRNA fragments from the polyacrylamide gel is sufficient.
For RNA-seq, mRNA is obtained by the depletion of rRNA from the total RNA - rRNA molecules hybridize to the biotinylated oligonucleotide probes which bind to the streptavidin-coated magnetic beads. The rRNA-oligonucleotide-bead complexes are then removed from the sample with a magnet resulting in a rRNA depleted sample19,20. The purified mRNA molecules are then randomly fragmented by alkaline hydrolysis. The obtained fragments of mRNA as well as the ribosomal footprints are converted into cDNA libraries and prepared for deep sequencing (Figure 2). This involves ends repair needed after alkaline hydrolysis (for mRNA) and enzymatic digestion (for RF): dephosphorylation of 3&#39; ends followed by phosphorylation of 5&#39; ends. The next steps are adaptors ligation and the reverse transcription to create cDNA inserts framed by sequences required for the next generation sequencing (NGS) using Illumina platform. The last phase of library preparation is a PCR reaction in which the constructs are amplified and labelled with sample specific barcodes to allow multiplexing and sequencing various samples on one channel. Before sequencing, the quality and quantity of the libraries are assessed by the high-sensitivity DNA on-chip electrophoresis. cDNA libraries with appropriate parameters can then be pooled and sequenced. Sequencing can be performed on different Illumina platforms, such as MiSeq, NextSeq or HighSeq, depending on the number of libraries, required read length and sequencing depth. After sequencing, the bioinformatic analysis is performed.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62544/62544fig02.jpg" /> 
Figure 2. Library preparation. Library preparation includes the ends repair, adapters ligation, reverse transcription and amplification with barcoding. <a href="https://www.jove.com/files/ftp_upload/62544/62544fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The ribosome profiling is a universal method which can be easily modified and adjusted according to the scientific question. Originally it was used in yeast1, but shortly after it was applied to bacterial cells21 as well as eukaryotic model organisms including mouse10, zebrafish22, fruit fly23 and Arabidopsis thaliana24. It was also used for studying different ribosome types: cytoplasmic, mitochondrial25,26 and chloroplast27,28. In eukaryotes RIBO-seq is commonly adapted and refined to investigate specific aspects of translation, including initiation10,11,29,30,31,32, elongation1,10,11,31,33, ribosome stalling33 and conformation change33. Most of the modifications involve the use of different translation inhibitors. In bacteria however, analogous studies have been difficult to conduct because of the paucity of inhibitors with the required mechanism of action34. The most commonly used translation inhibitor in bacteria is chloramphenicol (CAM) which binds to the peptidyl transferase center (PTC) and prevents correct positioning of the aminoacyl-tRNA in the A-site. As a result, CAM prevents the formation of a peptide bond which leads to arresting the elongating ribosomes35. Other examples of translation inhibitors in bacteria are tetracycline (TET)36, retapamulin (RET)34 and Onc11237 which have been used to investigate translation initiation sites. TET, which prevents tRNA delivery to the ribosome by directly overlapping with the anticodon stem-loop of tRNA at the A-site, was originally applied to verify the results obtained from CAM treatment since they are both antibiotics inhibiting translation elongation38. TET was found to detect primary TIS, however was unable to reveal internal TIS36. RET binds in the PTC of the bacterial ribosome, and prevents formation of the first peptide bond by interfering with an elongator aminoacyl-tRNA in the A site. Applying RET results in ribosomes arrest at both primary as well as internal TISs34. Onc112, a proline-rich antimicrobial peptide, binds in the exit tunnel and blocks aminoacyl-tRNA binding in the ribosomal A site. As a result, Onc112 prevents initiation complexes from entering the elongation phase37.&#160;
The main information ribosome profiling provides is ribosomes density and their position on the mRNA. It was successfully applied to investigate differential gene expression at the level of translation in various growth conditions1,6, measure translational efficiency1,38,39 and detect translation regulation events such as ribosomal pausing10. RIBO-seq also allows for uncovering the translation of annotated ncRNA, pseudogenes and unannotated small open reading frames (ORF) leading to the identification of novel and/or very short protein coding genes10,12,22,30,37. In such cases, RIBO-seq can fine-tune and improve genome annotation. With its high sensitivity for the identification of translated ORFs and its quantitative nature, ribosome profiling can also serve as a proxy for the proteome determination or in aiding proteomics studies31,34,39. By mapping TIS, ribosome profiling reveals N-terminally extended and truncated isoforms of known proteins10,32. RIBO-seq was also adapted to study co-translational folding of proteins14,21,24. This&#160;method enables measuring of elongation rates1,10,39 or decoding speeds of individual codons6 and helps in developing quantitative models of translation17. The ribosome profiling method is also capable of providing mechanistic insights into the ribosome pausing in bacteria7,15,17, frameshifting40, stop-codon readthrough21, termination/recycling defects41,42 and ribosomal conformation changes33 in eukaryotes. RIBO-seq was also adapted to examine the impact of specific trans-acting factors on translation such as miRNAs6 and RNA-binding proteins in eukaryotes16,43. However, it is important to acknowledge that the experimental design and the obtained resolution of RIBO-seq determine the amount of information that can be extracted from the resulting sequencing data12.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>10X TBE (powder)</td>
			<td>Invitrogen</td>
			<td>AM9864</td>
			<td></td>
		</tr>
		<tr>
			<td>2-Mercaptoethanol, 99%, pure</td>
			<td>Acros Organics</td>
			<td>125472500</td>
			<td></td>
		</tr>
		<tr>
			<td>Adenosine 5&#39;-Triphosphate (ATP)</td>
			<td>New England Biolabs</td>
			<td>P0756S</td>
			<td></td>
		</tr>
		<tr>
			<td>Aluminium oxide calcinated pure p.a.</td>
			<td>Chempur</td>
			<td>114560600</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium chloride dihydrate</td>
			<td>Sigma-Aldrich</td>
			<td>C3881-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Chloramphenicol</td>
			<td>MP Biomedicals</td>
			<td>190321</td>
			<td></td>
		</tr>
		<tr>
			<td>DNA Clean &#38; Concentrator -5</td>
			<td>Zymo Research</td>
			<td>D4004</td>
			<td></td>
		</tr>
		<tr>
			<td>Dnase I recombinant, Rnase-free</td>
			<td>Roche</td>
			<td>4716728001</td>
			<td></td>
		</tr>
		<tr>
			<td>EDTA disodium salt</td>
			<td>Fisher Scientific</td>
			<td>E/P140/48</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethyl Alcohol Absolut 99,8%&#160; Pure-P.A.-Basic</td>
			<td>POCH Avantor Performance Materials Poland S.A</td>
			<td>BA6480111</td>
			<td></td>
		</tr>
		<tr>
			<td>Filtration apparatus</td>
			<td>VWR Collection</td>
			<td>511-0265</td>
			<td>all-glass filtration apparatus, with funnel, fritted base, cap, 47 mm &#216; spring clamp and ground joint flask</td>
		</tr>
		<tr>
			<td>Gel 40 (19:1)</td>
			<td>Rotiphorese</td>
			<td>3030.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Gel Loading Dye, Blue, 6X</td>
			<td>New England Biolabs</td>
			<td>E6138G</td>
			<td></td>
		</tr>
		<tr>
			<td>Guanosine 5&#8242;-[&#946;,&#947;-imido]triphosphate trisodium salt hydrate</td>
			<td>Sigma-Aldrich</td>
			<td>G0635-25MG</td>
			<td></td>
		</tr>
		<tr>
			<td>labZAP</td>
			<td>A&#38;A Biotechnology</td>
			<td>040-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium acetate tetrahydrate</td>
			<td>Sigma-Aldrich</td>
			<td>M5661-250G</td>
			<td></td>
		</tr>
		<tr>
			<td>MCE membrane fiter</td>
			<td>Alfatec Technology</td>
			<td>M47MCE45GWS</td>
			<td>pore size: 0.45um</td>
		</tr>
		<tr>
			<td>MICROBExpress Bacterial mRNA Purification</td>
			<td>Invitrogen</td>
			<td>AM1905</td>
			<td></td>
		</tr>
		<tr>
			<td>Multiplex Small RNA Library Prep Set for Illumina</td>
			<td>New England Biolabs</td>
			<td>E7300S</td>
			<td></td>
		</tr>
		<tr>
			<td>Nuclease-Free Water</td>
			<td>Ambion</td>
			<td>AM9937</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium Acetate Anhydrous Pure P.A.</td>
			<td>POCH Avantor Performance Materials Poland S.A</td>
			<td>744330113</td>
			<td></td>
		</tr>
		<tr>
			<td>Quick-Load pBR322 DNA-MspI Digest</td>
			<td>New England Biolabs</td>
			<td>E7323A</td>
			<td></td>
		</tr>
		<tr>
			<td>RNA Clean &#38; Concentrator -25</td>
			<td>Zymo Research</td>
			<td>R1018</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium acetate</td>
			<td>Sigma-Aldrich</td>
			<td>S2889-250G</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium carbonate</td>
			<td>Sigma-Aldrich</td>
			<td>223530-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium hydrogen carbonate pure p.a.</td>
			<td>POCH Avantor Performance Materials Poland S.A</td>
			<td>810530115</td>
			<td></td>
		</tr>
		<tr>
			<td>SYBR Gold nucleic acid gel stain</td>
			<td>Life Technologies</td>
			<td>S11494</td>
			<td></td>
		</tr>
		<tr>
			<td>T4 Polynucleotide Kinase</td>
			<td>New England Biolabs</td>
			<td>M0201L</td>
			<td></td>
		</tr>
		<tr>
			<td>T4 Polynucleotide Kinase Reaction Buffer</td>
			<td>New England Biolabs</td>
			<td>B0201S</td>
			<td></td>
		</tr>
		<tr>
			<td>TBE-Urea Sample Buffer (2x)</td>
			<td>Invitrogen</td>
			<td>LC6876</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris(hydroxymethyl)amino-methane, ultrapure, 99,9%</td>
			<td>AlfaAesar</td>
			<td>J65594</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100, 98%</td>
			<td>Acros Organics</td>
			<td>327371000</td>
			<td></td>
		</tr>
		<tr>
			<td>Urea G.R.</td>
			<td>lach:ner</td>
			<td>40096-AP0</td>
			<td></td>
		</tr>
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ribo-seq in bacteria: a sample collection and library preparation protocol for ngs sequencing

The ribosome profiling technique (RIBO-seq) is currently the most effective tool for studying the process of protein synthesis in vivo. The advantage of this method, in comparison to other approaches, is its ability to monitor translation by precisely mapping the position and number of ribosomes on a mRNA transcript.
In this article, we describe the consecutive stages of sample collection and preparation for RIBO-seq method in bacteria, highlighting the details relevant to the planning and execution of the experiment.
Since the RIBO-seq relies on intact ribosomes and related mRNAs, the key step is rapid inhibition of translation and adequate disintegration of cells. Thus, we suggest filtration and flash-freezing in liquid nitrogen for cell harvesting with an optional pretreatment with chloramphenicol to arrest translation in bacteria. For the disintegration, we propose grinding frozen cells with mortar and pestle in the presence of aluminum oxide to mechanically disrupt the cell wall. In this protocol, sucrose cushion or a sucrose gradient ultracentrifugation for monosome purification is not required. Instead, mRNA separation using polyacrylamide gel electrophoresis (PAGE) followed by the ribosomal footprint excision (28-30 nt band) is applied and provides satisfactory results. This largely simplifies the method as well as reduces the time and equipment requirements for the procedure. For library preparation, we recommend using the commercially available small RNA kit for Illumina sequencing from New England Biolabs, following manufacturer&#39;s guidelines with some degree of optimization.
The resulting cDNA libraries present appropriate quantity and quality required for next generation sequencing (NGS). Sequencing of the libraries prepared according to the described protocol results in 2 to 10 mln uniquely mapped reads per sample providing sufficient data for comprehensive bioinformatic analysis. The protocol we present is quick and relatively easy and can be performed with standard laboratory equipment.

The exemplary results presented here were obtained in a study examining translation regulation in sporulating WT Bacillus subtilis cells. Overnight cultures were diluted to OD600 equal to 0.1 in 100 mL of rich medium and incubated at 37 &#176;C with vigorous shaking until OD600 reached 0.5-0.6. The rich medium was then replaced with minimal medium to induce sporulation process and the incubation was continued for up to four hours. Cells were harvested every hour beginning with T0 - sporulat...

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RIBO-seq in Bacteria: a Sample Collection and Library Preparation Protocol for NGS Sequencing

Here we describe the stages of sample collection and preparation for RIBO-seq in bacteria. Sequencing of the libraries prepared according to these guidelines results in sufficient data for comprehensive bioinformatic analysis. The protocol we present is simple, uses standard laboratory equipment and takes seven days from lysis to obtaining libraries.

The ribosome profiling technique (RIBO-seq) is currently the most effective tool for studying the process of protein synthesis in vivo. The advantage of ...

ribo-seq-bacteria-sample-collection-library-preparation-protocol-for

Research

JoVE Journal

Biochemistry

8.0K Views.  Maria Curie-Sklodowska University. Here we describe the stages of sample collection and preparation for RIBO-seq in bacteria. Sequencing of the libraries prepared according to these guidelines results in sufficient data for comprehensive bioinformatic analysis. The protocol we present is simple, uses standard laboratory equipment and takes seven days from lysis to obtaining libraries. 

Video: RIBO-seq in Bacteria: a Sample Collection and Library Preparation Protocol for NGS Sequencing

Sample Preparation for Mass Spectrometry-based Identification of RNA-binding Regions

Noncoding RNAs play important roles in several nuclear processes, including regulating gene expression, chromatin structure, and DNA repair. In most cases, the action of noncoding RNAs is mediated by proteins whose functions are in turn regulated by these interactions with noncoding RNAs. Consistent with this, a growing number of proteins involved in nuclear functions have been reported to bind RNA and in a few cases the RNA-binding regions of these proteins have been mapped, often through laborious, candidate-based methods.
Here, we report a detailed protocol to perform a high-throughput, proteome-wide unbiased identification of RNA-binding proteins and their RNA-binding regions. The methodology relies on the incorporation of a photoreactive uridine analog in the cellular RNA, followed by UV-mediated protein-RNA crosslinking, and mass spectrometry analyses to reveal RNA-crosslinked peptides within the proteome. Although we describe the procedure for mouse embryonic stem cells, the protocol should be easily adapted to a variety of cultured cells.

We describe a protocol to identify RNA-binding proteins and map their RNA-binding regions in live cells using UV-mediated photocrosslinking and mass spectrometry.

Noncoding RNAs play important roles in several nuclear processes, including regulating gene expression, chromatin structure, and DNA repair. In most ...

An All-in-one Sample Holder for Macromolecular X-ray Crystallography with Minimal Background Scattering

Macromolecular X-ray crystallography (MX) is the most prominent method to obtain high-resolution three-dimensional knowledge of biological macromolecules. A prerequisite for the method is that highly ordered crystalline specimen need to be grown from the macromolecule to be studied, which then need to be prepared for the diffraction experiment. This preparation procedure typically involves removal of the crystal from the solution, in which it was grown, soaking of the crystal in ligand solution or cryo-protectant solution and then immobilizing the crystal on a mount suitable for the experiment. A serious problem for this procedure is that macromolecular crystals are often mechanically unstable and rather fragile. Consequently, the handling of such fragile crystals can easily become a bottleneck in a structure determination attempt. Any mechanical force applied to such delicate crystals may disturb the regular packing of the molecules and may lead to a loss of diffraction power of the crystals. Here, we present a novel all-in-one sample holder, which has been developed in order to minimize the handling steps of crystals and hence to maximize the success rate of the structure determination experiment. The sample holder supports the setup of crystal drops by replacing the commonly used microscope cover slips. Further, it allows in-place crystal manipulation such as ligand soaking, cryo-protection and complex formation without any opening of the crystallization cavity and without crystal handling. Finally, the sample holder has been designed in order to enable the collection of in situ X-ray diffraction data at both, ambient and cryogenic temperature. By using this sample holder, the chances to damage the crystal on its way from crystallization to diffraction data collection are considerably reduced since direct crystal handling is no longer required.

A novel sample holder for macromolecular X-ray crystallography along with a suitable handling protocol is presented. The system allows crystal growth, crystal soaking and in situ diffraction data collection at both, ambient and cryogenic temperature without the need of any crystal manipulation or mounting.

Macromolecular X-ray crystallography (MX) is the most prominent method to obtain high-resolution three-dimensional knowledge of biological macromolecules. ...

A Plasma Sample Preparation for Mass Spectrometry using an Automated Workstation

Sample preparation for mass spectrometry analysis in proteomics requires enzymatic cleavage of proteins into a peptide mixture. This process involves numerous incubation and liquid transfer steps in order to achieve denaturation, reduction, alkylation, and cleavage. Adapting this workflow onto an automated workstation can increase efficiency and reduce coefficients of variance, thereby providing more reliable data for statistical comparisons between sample types. We previously described an automated proteomic sample preparation workflow1. Here, we report the development of a more efficient and better controlled workflow with the following advantages: 1) The number of liquid transfer steps is reduced from nine to six by combining reagents; 2) Pipetting time is reduced by selective tip pipetting using a 96-position pipetting head with multiple channels; 3) Potential throughput is increased by the availability of up to 45 deck positions; 4) Complete enclosure of the system provides improved temperature and environmental control and reduces the potential for contamination of samples or reagents; and 5) The addition of stable isotope labeled peptides, as well as &#946;-galactosidase protein, to each sample makes monitoring and quality control possible throughout the entire process. These hardware and process improvements provide good reproducibility and improve intra-assay and inter-assay precision (CV of less than 20%) for LC-MS based protein and peptide quantification. The entire workflow for digesting 96 samples in a 96-well plate can be completed in approximately 5 hours.

Here, we present an automated plasma protein digestion method for mass spectrometry-based quantitative proteomic analysis. In this protocol, the liquid transferring and incubation steps for protein denaturation, reduction, alkylation, and trypsin digestion reactions are streamlined and automated. It takes approximately five hours to prepare a 96-well plate with desired precision.

Sample preparation for mass spectrometry analysis in proteomics requires enzymatic cleavage of proteins into a peptide mixture. This process involves ...

Sample Preparation for Probe Electrospray Ionization Mass Spectrometry

Mass spectrometry (MS) is a powerful tool in analytical chemistry because it provides very accurate information about molecules, such as mass-to-charge ratios (m/z), which are useful to deduce molecular weights and structures. While it is essentially a destructive analytical method, recent advancements in the ambient ionization technique have enabled us to acquire data while leaving tissue in a relatively intact state in terms of integrity. Probe electrospray ionization (PESI) is a so-called direct method because it does not require complex and time-consuming pretreatment of samples. A fine needle serves as a sample picker, as well as an ionization emitter. Based on the very sharp and fine property of the probe tip, destruction of the samples is minimal, allowing us to acquire the real-time molecular information from living things in situ. Herein, we introduce three applications of PESI-MS technique that will be useful for biomedical research and development. One involves the application to solid tissue, which is the basic application of this technique for the medical diagnosis. As this technique requires only 10 mg of the sample, it may be very useful in the routine clinical settings. The second application is for in vitro medical diagnostics where human blood serum is measured. The ability to measure fluid samples is also valuable in various biological experiments where a sufficient volume of sample for conventional analytical techniques cannot be provided. The third application leans toward the direct application of probe needles in living animals, where we can obtain real-time dynamics of metabolites or drugs in specific organs. In each application, we can infer the molecules that have been detected by MS or use artificial intelligence to obtain a medical diagnosis.

This article introduces sample preparation methods for a unique real-time analytical method based on the ambient mass spectrometry. This method lets us perform real-time analysis of the biological molecules in vivo without any special pretreatments.

Mass spectrometry (MS) is a powerful tool in analytical chemistry because it provides very accurate information about molecules, such as mass-to-charge ...

TMT Sample Preparation for Proteomics Facility Submission and Subsequent Data Analysis

Proteomic technologies are powerful methodologies that can aid our understanding of mechanisms of action in biological systems by providing a global view of the impact of a disease, treatment, or other condition on the proteome as a whole. This report provides a detailed protocol for the extraction, quantification, precipitation, digestion, labeling, and subsequent data analysis of protein samples. Our optimized TMT labeling protocol requires a lower tag-label concentration and achieves consistently reliable data. We have used this protocol to evaluate protein expression profiles in a variety of mouse tissues (i.e., heart, skeletal muscle, and brain) as well as cells cultured in vitro. In addition, we demonstrate how to evaluate thousands of proteins from the resulting dataset.

We present an optimized tandem mass tag (TMT) labeling protocol that includes detailed information for each of the following steps: protein extraction, quantification, precipitation, digestion, labeling, submission to a proteomics facility, and data analyses.

Proteomic technologies are powerful methodologies that can aid our understanding of mechanisms of action in biological systems by providing a global view ...

MS2-Affinity Purification Coupled with RNA Sequencing in Gram-Positive Bacteria

Although small regulatory RNAs (sRNAs) are widespread among the bacterial domain of life, the functions of many of them remain poorly characterized notably due to the difficulty of identifying their mRNA targets. Here, we described a modified protocol of the MS2-Affinity Purification coupled with RNA Sequencing (MAPS) technology, aiming to reveal all RNA partners of a specific sRNA in vivo. Broadly, the MS2 aptamer is fused to the 5&#8217; extremity of the sRNA of interest. This construct is then expressed in vivo, allowing the MS2-sRNA to interact with its cellular partners. After bacterial harvesting, cells are mechanically lysed. The crude extract is loaded into an amylose-based chromatography column previously coated with the MS2 protein fused to the maltose binding protein. This enables the specific capture of MS2-sRNA and interacting RNAs. After elution, co-purified RNAs are identified by high-throughput RNA sequencing and subsequent bioinformatic analysis. The following protocol has been implemented in the Gram-positive human pathogen Staphylococcus aureus and is, in principle, transposable to any Gram-positive bacteria. To sum up, MAPS technology constitutes an efficient method to deeply explore the regulatory network of a particular sRNA, offering a snapshot of its whole targetome. However, it is important to keep in mind that putative targets identified by MAPS still need to be validated by complementary experimental approaches.

MAPS technology has been developed to scrutinize the targetome of a specific regulatory RNA in vivo. The sRNA of interest is tagged with a MS2 aptamer enabling the co-purification of its RNA partners and their identification by RNA sequencing. This modified protocol is particularly suited for Gram-positive bacteria.&#160;

Although small regulatory RNAs (sRNAs) are widespread among the bacterial domain of life, the functions of many of them remain poorly characterized ...

Plant Sample Preparation for Nucleoside/Nucleotide Content Measurement with An HPLC-MS/MS

Nucleosides/nucleotides are building blocks of nucleic acids, parts of cosubstrates and coenzymes, cell signaling molecules, and energy carriers, which are involved in many cell activities. Here, we describe a rapid and reliable method for the absolute qualification of nucleoside/nucleotide contents in plants. Briefly, 100 mg of homogenized plant material was extracted with 1 mL of extraction buffer (methanol, acetonitrile, and water at a ratio of 2:2:1). Later, the sample was concentrated five times in a freeze dryer and then injected into an HPLC-MS/MS. Nucleotides were separated on a porous graphitic carbon (PGC) column and nucleosides were separated on a C18 column. The mass transitions of each nucleoside and nucleotide were monitored by mass spectrometry. The contents of the nucleosides and nucleotides were quantified against their external standards (ESTDs). Using this method, therefore, researchers can easily quantify nucleosides/nucleotides in different plants.

A precise and reproducible method for in vivo nucleosides/nucleotides quantification in plants is described here. This method employs an HPLC-MS/MS.

Nucleosides/nucleotides are building blocks of nucleic acids, parts of cosubstrates and coenzymes, cell signaling molecules, and energy carriers, which ...

NMR-Based Fragment Screening in a Minimum Sample but Maximum Automation Mode

Fragment-based screening (FBS) is a well-validated and accepted concept within the drug discovery process both in academia and industry. The greatest advantage of NMR-based fragment screening is its ability not only to detect binders over 7-8 orders of magnitude of affinity but also to monitor purity and chemical quality of the fragments and thus to produce high quality hits and minimal false positives or false negatives. A prerequisite within the FBS is to perform initial and periodic quality control of the fragment library, determining solubility and chemical integrity of the fragments in relevant buffers, and establishing multiple libraries to cover diverse scaffolds to accommodate various macromolecule target classes (proteins/RNA/DNA). Further, an extensive NMR-based screening protocol optimization with respect to sample quantities, speed of acquisition and analysis at the level of biological construct/fragment-space, in condition-space (buffer, additives, ions, pH, and temperature) and in ligand-space (ligand analogues, ligand concentration) is required. At least in academia, these screening efforts have so far been undertaken manually in a very limited fashion, leading to limited availability of screening infrastructure not only in the drug development process but also in the context of chemical probe development. In order to meet the requirements economically, advanced workflows are presented. They take advantage of the latest state-of-the-art advanced hardware, with which the liquid sample collection can be filled in a temperature-controlled fashion into the NMR-tubes in an automated manner. 1H/19F NMR ligand-based spectra are then collected at a given temperature. High-throughput sample changer (HT sample changer) can handle more than 500 samples in temperature-controlled blocks. This together with advanced software tools speeds up data acquisition and analysis. Further, application of screening routines on protein and RNA samples are described to make aware of the established protocols for a broad user base in biomacromolecular research.

Fragment-based screening by NMR is a robust method to rapidly identify small molecule binders to biomacromolecules (DNA, RNA, or proteins). Protocols describing automation-based sample preparation, NMR experiments &#38; acquisition conditions, and analysis workflows are presented. The technique allows for optimal exploitation of both 1H and 19F NMR-active nuclei for detection.

Fragment-based screening (FBS) is a well-validated and accepted concept within the drug discovery process both in academia and industry. The greatest ...

A Sample Preparation Pipeline for Microcrystals at the VMXm Beamline

The mounting of microcrystals (&#60;10 &#181;m) for single crystal cryo-crystallography presents a non-trivial challenge. Improvements in data quality have been seen for microcrystals with the development of beamline optics, beam stability and variable beam size focusing from submicron to microns, such as at the VMXm beamline at Diamond Light Source1. Further improvements in data quality will be gained through improvements in sample environment and sample preparation. Microcrystals inherently generate weaker diffraction, therefore improving the signal-to-noise is key to collecting quality X-ray diffraction data and will predominantly come from reductions in background noise. Major sources of X-ray background noise in a diffraction experiment are from their interaction with the air path before and after the sample, excess crystallization solution surrounding the sample, the presence of crystalline ice and scatter from any other beamline instrumentation or X-ray windows. The VMXm beamline comprises instrumentation and a sample preparation protocol to reduce all these sources of noise.
Firstly, an in-vacuum sample environment at VMXm removes the air path between X-ray source and sample. Next, sample preparation protocols for macromolecular crystallography at VMXm utilize a number of processes and tools adapted from cryoTEM. These include copper grids with holey carbon support films, automated blotting and plunge cooling robotics making use of liquid ethane. These tools enable the preparation of hundreds of microcrystals on a single cryoTEM grid with minimal surrounding liquid on a low-noise support. They also minimize the formation of crystalline ice from any remaining liquid surrounding the crystals. 
We present the process for preparing and assessing the quality of soluble protein microcrystals using visible light and scanning electron microscopy before mounting the samples on the VMXm beamline for X-ray diffraction experiments. We will also provide examples of good quality samples as well as those which require further optimization and strategies to do so.

The signal-to-noise ratio of data is one of the most important considerations in performing X-ray diffraction measurements from microcrystals. The VMXm beamline provides a low-noise environment and microbeam for such experiments. Here, we describe sample preparation methods for mounting and cooling microcrystals for VMXm and other microfocus macromolecular crystallography beamlines.

The mounting of microcrystals (&#60;10 &#181;m) for single crystal cryo-crystallography presents a non-trivial challenge. Improvements in data quality ...

Sample Preparation and Transfer Protocol for In-Vacuum Long-Wavelength Crystallography on Beamline I23 at Diamond Light Source

Long-wavelength macromolecular crystallography (MX) exploits the anomalous scattering properties of elements, such as sulfur, phosphorus, potassium, chlorine, or calcium, that are often natively present in macromolecules. This enables the direct structure solution of proteins and nucleic acids via experimental phasing without the need of additional labelling. To eliminate the significant air absorption of X-rays in this wavelength regime, these experiments are performed in a vacuum environment. Beamline I23 at Diamond Light Source, UK, is the first synchrotron instrument of its kind, designed and optimized for MX experiments in the long wavelength range towards 5 &#197;. 
To make this possible, a large vacuum vessel encloses all endstation components of the sample environment. The necessity to maintain samples at cryogenic temperatures during storage and data collection in vacuum requires the use of thermally conductive sample holders. This facilitates efficient heat removal to ensure sample cooling to approximately 50 K. The current protocol describes the procedures used for sample preparation and transfer of samples into vacuum on beamline I23. Ensuring uniformity in practices and methods already established within the macromolecular crystallography community, sample cooling to liquid nitrogen temperature can be performed in any laboratory setting equipped with standard MX tools. 
Cryogenic storage and transport of samples only require standard commercially available equipment. Specialized equipment is required for the transfer of cryogenically cooled crystals from liquid nitrogen into the vacuum endstation. Bespoke sample handling tools and a dedicated Cryogenic Transfer System (CTS) have been developed in house. Diffraction data collected on samples prepared using this protocol show excellent merging statistics, indicating that the quality of samples is unaltered during the procedure. This opens unique opportunities for in-vacuum MX in a wavelength range beyond standard synchrotron beamlines.

Here, we present a protocol for cryogenic sample preparation and transfer of crystals into the vacuum endstation on beamline I23 at Diamond Light Source, for long-wavelength macromolecular X-ray crystallography experiments.

Long-wavelength macromolecular crystallography (MX) exploits the anomalous scattering properties of elements, such as sulfur, phosphorus, potassium, ...