Ribosome Profiling in the Model Diatom Thalassiosira pseudonana

Diatoms are an important group of eukaryotic microalgae, which play key roles in marine biochemical cycling and possess significant biotechnological potential. Despite the importance of diatoms, their regulatory mechanisms of protein synthesis at the translational level remain largely unexplored. Here, we describe the detailed development of a ribosome profiling protocol to study translation in the model diatom Thalassiosira pseudonana, which can easily be adopted for other diatom species. To isolate and sequence ribosome-protected mRNA, total RNA was digested, and the ribosome-protected fragments were obtained by a combination of sucrose-cushion ultracentrifugation and polyacrylamide gel electrophoresis for size selection. To minimize rRNA contamination, a subtractive hybridization step using biotinylated oligos was employed. Subsequently, fragments were converted into sequencing libraries, enabling the global quantification and analysis of changes in protein synthesis in diatoms. The development of this novel ribosome profiling protocol represents a major expansion of the molecular toolbox available for diatoms and therefore has the potential to advance our understanding of the translational regulation in this important group of phytoplankton. © 2023 The Authors. Current Protocols published by Wiley Periodicals LLC. Basic Protocol: Ribosome profiling in Thalassiosira pseudonana Alternate Protocol

Diatoms are an important group of eukaryotic microalgae, which play key roles in marine biochemical cycling and possess significant biotechnological potential. Despite the importance of diatoms, their regulatory mechanisms of protein synthesis at the translational level remain largely unexplored. Here, we describe the detailed development of a ribosome profiling protocol to study translation in the model diatom Thalassiosira pseudonana, which can easily be adopted for other diatom species. To isolate and sequence ribosome-protected mRNA, total RNA was digested, and the ribosome-protected fragments were obtained by a combination of sucrose-cushion ultracentrifugation and polyacrylamide gel electrophoresis for size selection. To minimize rRNA contamination, a subtractive hybridization step using biotinylated oligos was employed. Subsequently, fragments were converted into sequencing libraries, enabling the global quantification and analysis of changes in protein synthesis in diatoms. The development of this novel ribosome profiling protocol represents a major expansion of the molecular toolbox available for diatoms and therefore has the potential to advance our understanding of the translational regulation in this important group of phytoplankton. © 2023 The Authors. Current Protocols published by Wiley Periodicals LLC.

INTRODUCTION
Diatoms are unicellular, eukaryotic microalgae, which comprise over 100,000 species throughout all aquatic environments. Their mosaic genomes have been shaped by secondary endosymbiosis and horizontal gene transfer, providing them with diverse regulatory mechanisms to adapt their protein synthesis in response to the changing environmental conditions (Mock et al., 2022). Due to their key role as primary producers in aquatic systems and their biotechnological potential, gene regulatory mechanisms in diatoms have been extensively studied using transcriptomic and proteomic data (e.g., Dong et al., 2016;Mock et al., 2008). The globally distributed species Thalassiosira pseudonana CCMP1335 was chosen for the first diatom genome sequencing project (Armbrust et al., 2004). Subsequently, it became a model organism due to the development of genetic manipulation tools such as the incorporation of recombinant DNA via transformation (Poulsen et al., 2006) and CRISPR/Cas-based genome editing (e.g., Belshaw et al., 2023;Hopes et al., 2016). However, despite these developments, regulation of protein synthesis at the translational level is largely unexplored in T. pseudonana and diatoms in general. The development of ribosome profiling or Ribo-Seq in 2009 (Ingolia et al., 2009) spurred the transcriptome-wide monitoring and quantification of translation in diatoms. This technique is based on the analysis of ∼30 nucleotide (nt) long mRNA fragments which are enclosed by translating ribosomes and protected from nuclease digestion (Steitz, 1969) (Fig. 1A). Deep sequencing of these generated footprints or ribosomeprotected fragments (RPFs) thus provides a genome-wide and high-resolution snapshot of translation (Ingolia et al., 2009). Ribosome profiling was first developed in Saccharomyces cerevisiae (Ingolia et al., 2009) and has since been successfully applied to study translation in a wide range of organisms, such as bacteria (Li et al., 2012), fishes (Bazzini et al., 2012), mammals (Guo et al., 2010;Ingolia et al., 2011) and plants (Hsu et al., 2016;Juntawong et al., 2014). Ribosome profiling has revolutionized our understanding of gene expression by unveiling the full complexity and regulation of translation in both prokaryotes and eukaryotes. It has provided novel mechanistic insights into the translation mechanism, such as ribosomal pausing sites (Brar & Weissman, 2015;Ingolia et al., 2011;Karlsen et al., 2018;Shalgi et al., 2013) or previously unannotated (small) open reading frames (ORFs) (Aspden et al., 2014;Bazzini et al., 2014;Calviello et al., 2016;Hsu et al., 2016;Ji et al., 2015;Wu et al., 2019). However, to date, ribosome profiling has only been performed on one algal species, the green alga Chlamydomonas reinhardtii (Chung et al., 2015;Trosch et al., 2018). Here, we report the development of a ribosome profiling protocol for the model diatom T. pseudonana CCMP1335, representing the first application of this technique for any marine algae. Previously developed protocols were adapted (McGlincy & Ingolia, 2017;Meindl et al., 2023;, which involved optimizations of harvesting strategy and cell lysis conditions, the amount of nuclease for mRNA digestion, and the implementation of a subtractive hybridization step for rRNA removal. Our optimized strategy ensures the generation of high-quality footprint data for this important class of organisms. Thus, the application of ribosome profiling in diatoms provides a powerful tool for studying their translational regulation. Moreover, Figure 1 (A) Overview of the ribosome profiling protocol. The ribosome profiling protocol for T. pseudonana follows the same basic steps as for other cell types. Cells are harvested and lysed before nuclease digestion of mRNA unprotected by ribosomes is performed. mRNA fragments bound by ribosomes are recovered and purified prior to library preparation, sequencing, and computational data analysis. (B) Workflow diagram depicting all steps used in our ribosome profiling protocol. The four main parts include: (1) lysate preparation, (2) generation of ribosome protected footprints and footprint purification, (3) library preparation, and (4) sequencing and data processing. The corresponding steps in the Basic Protocol are listed. Additional steps outlined in the Alternate Protocol are necessary if sucrose density fractionation is performed. this protocol can be adapted for use with other diatom species and may also facilitate the study of other marine algae.
The Basic Protocol involves a series of steps for monosome preparation, isolation of ribosome-protected fragments, and subsequent sequencing library preparation (Fig. 1B). We have also developed an Alternate Protocol that yields comparable results but utilizes a sucrose gradient instead of a sucrose cushion, therefore requiring additional instrumentation. This Alternate Protocol enables quality control of the samples prior to sequencing library preparation, facilitating the adaption of ribosome profiling to other diatom species, which typically requires the optimization of reaction conditions, such as those used for limited nucleolytic digestion.

STRATEGIC PLANNING
To ensure optimal results, ribosome profiling should be carried out using exponentially growing T. pseudonana cells. For all experiments, we recommend a minimum of three biological replicates per condition to enhance data robustness. Additionally, to limit ribosomal run-off and RNA degradation, several steps of the protocol must be carried out in a cold room or on ice. As this protocol utilizes potentially hazardous chemicals such as cycloheximide and organic solvents, careful handling and access to a fume hood is crucial. Following monosome purification, it is imperative to maintain an RNase-free environment, using dedicated workspaces and RNase-free reagents. To minimize variation due to batch effects, it is advised to process samples from different conditions together by the Pichler et al.

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RIBOSOME PROFILING IN THALASSIOSIRA PSEUDONANA
The following protocol is designed to generate high-quality ribosome profiling data to quantify translation in T. pseudonana CCMP1335. Initially, cells are promptly harvested and flash-frozen in liquid nitrogen. After mechanical disruption of the cells, optimized RNase digestion conditions are employed to create ribosome-protected footprints. Monosomes are subsequently isolated using sucrose-cushion ultracentrifugation. Alternatively, ribosomes can be purified via sucrose density gradient fractionation as detailed in the Alternate Protocol. This protocol typically yields ∼10 to 50 ng of 26 to 31 nt RNA fragments (corresponding to ∼1 to 5 pmol of RNA), which serves as the optimal amount of starting material for sequencing library preparation. We have incorporated an rRNA removal step via subtractive hybridization with the aim to increase the fraction of informative reads that map to mRNAs.

Materials
Thalassiosira pseudonana clone CCMP1335 (Bigelow; see Internet Resources) Aquil medium (Price et al., 1989;  NI-800 and NI-801 size marker oligos (see Table 1) 10,000× SYBR Gold (ThermoFischer Scientific, cat. no. S11494) RNA gel extraction buffer (see recipe) Ethanol (Carl Roth, cat. no. 9065) Qubit microRNA assay kit (ThermoFisher Scientific, cat. no. Q32881) T4 polynucleotide kinase (PNK) and buffer (New England Biolabs, cat. no. M0201) RNaseOUT recombinant ribonuclease inhibitor (ThermoFisher Scientific, cat. no. 10777019) Pre-adenylated rApp-L7 (see Table 2) a Oligonucleotides used for library preparation are the same as described in Meindl et al. (2023). b All oligonucleotides are HPLC-purified. c Store and dilute primers in a buffered solution (e.g., 10 mM Tris·HCl, pH 8.0-8.5). To limit the number of freeze-thaw cycles, store as aliquots at −20°C. d 5' ribo-adenylated, 3' protected by di-deoxy nucleotide (ddC), contains a 3 nt 5' randomized sequence to minimize ligation bias and to serve as a UMI for the bioinformatic identification of PCR duplicates. e This RT oligonucleotide serves as a circularization adapter containing a short sequence on its 3' end that serves as a primer for reverse transcription. Separated by a PEG spacer (denoted iSp9 in the sequence) is a ligation adapter for the P7 side of the amplicon. It contains a 5' phosphate group followed by seven degenerate nucleotides that minimize ligation bias and that serve as UMIs for the bioinformatic identification of PCR duplicates. f To prevent degradation by the proof-reading activity of the polymerase, it is recommended to include a phosphorothioate bond at the 3' terminal end of the primer (highlighted as asterisk). g X marks the position of a 6 nt barcode for experimental multiplexing. See Internet Resources, Illumina TrueSeq single indexes.

Cell lysis
3. Add 400 μl of pre-chilled polysome resuspension buffer (PRB) to the tube with slightly thawed cells. Rinse off filter and remove with sterile tweezers. Add two scoops of glass beads (425-to 600-μm).
Lysis is done by bead beating of the frozen cell pellets. Prepare enough PRB (at least 1150 μl per tube) for lysis and sucrose cushion (step 10) using a 50-ml centrifuge tube.
4. Treat sample in bead beater for 2 min (max speed) with a break on ice after 1 min to prevent overheating.
5. Clarify the lysate by centrifugation for 10 min at 14,000 × g, 4°C using a microcentrifuge. Transfer the supernatant using a pipette in a new 1.5-ml microcentrifuge tube and place on ice.
6. Measure UV absorption at 260 nm of the lysate using a NanoDrop to estimate the RNA concentration. 11. Pellet ribosomes by centrifugation in an ultracentrifuge using a TLA120.2 rotor, 2 hr at 199,000 × g (75,000 rpm), 4°C.
While pelleted ribosomes are translucent, the pellet may also contain aggregates of the photosystems, which can aid in their identification. As a precaution, prior to removing the tubes from the rotor, it is recommended to mark their outside where the pelleted ribosomes are expected to be located.
12. Remove the supernatant and resuspend the pellet in 350 μl of TRizol reagent. Transfer to a 1.5-ml microcentrifuge tube. Replicates of the same sample can be pooled at this point.
13. Purify resuspended RNA by using the Direct-zol RNA MiniPrep kit following the manufacturer's instruction for purification of total RNA. Elute RNA in 50 μl RNase-free H 2 O provided with the kit.
15. Carry out precipitation on dry ice >30 min or at −20°C overnight.
16. Pellet RNA by centrifugation for 30 min at 14,000 × g, 4°C using a microcentrifuge. Pipette all liquid from the tube, place sideways and air dry for 10 min.
Samples can be stored overnight at −20˚C or for several months at −80°C.

Ribosomal footprint fragment purification
It is critical to work in an RNase-free environment from this step onwards. Decontaminate the electrophoresis apparatus and other equipment with RNaseZap. MilliQ H 2 O can be used to prepare the running buffer.
26. Visualize gel on a blue light transilluminator and cut out the 26 to 34 nt region of the gel using a scalpel blade (see Fig. 2A).

Gel extraction
The gel extraction step has been adapted from .
27. Using an 18-G needle, poke holes into a 0.5-ml tube. Place the gel cut out into the tube.
29. Spin 5 min at 14,000 × g, room temperature, in a microcentrifuge until gel extrudes into bottom tube.

Library preparation
For preparation of sequencing libraries, we followed the protocol by Meindl et al. (2023).
With this protocol sequencing libraries can be prepared from as little as 0.1 pmol of RNA fragments; an optimal amount of starting material is 1 to 5 pmol of RNA (from step 38).
To minimize sample loss, all reactions until PCR amplification of the final library (step 97) should be carried out in low bind reaction tubes.
We additionally implemented a subtractive hybridization step for rRNA removal using eight biotinylated depletion oligonucleotides representing abundant rRNA contaminants in T. pseudonana. The custom depletion step was optimized from Zinshteyn et al., and Green (2020).

RNA dephosphorylation to remove of terminal phosphates
40. For removal of the 2'-3'-cyclic phosphate that is generated by the RNase I cleavage, set up the following reaction:

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69. Incubate the mix for 5 min at room temperature. Magnetically attract the beads and add the eluate to the reverse transcription (RT) mix of the next step (step 70).

Reverse transcription
70. Prepare the following mix in 0.2-ml tubes: RNA and primer mix 12 μl pooled RNA in H 2 O (from step 69) 1 μ l 1 0m Md N T Pm i x 1 μl 0.5 pmol/μl RT Oligo P7-circ (see Table 2) 71. In a thermal cycler, heat the sample to 70°C for 5 min and incubate at 25°C until RT mix added, mix by pipetting. Do not put on ice to prevent unspecific annealing of the oligonucleotide used to prime reverse transcription.
72. Prepare the following RT mix and add it to the sample from the previous step (resulting in a total volume of 20 μl): 103. Repeat the last wash of the magnetically attracted beads with another 300 μl ProNex wash buffer for 40 to 60 s. Discard the supernatant and allow the samples to air-dry for ∼8 to 10 min (<60 min) until cracks are visible in the bead pellet.
104. Remove the beads from the magnetic stand and elute samples in 23 μl ProNex elution buffer. Resuspend all samples by pipetting and let them stand for 5 min at room temperature.
Samples can be stored at −20˚C for several months.

Second PCR amplification-cycle optimization
Try two different cycle numbers for amplification of the sample, as described in Buchbender et al. (2020). A good starting point for cycle optimization is a range of 6-12 cycles. Ideally, you should observe enough product without overamplification. Overamplification is indicated by the appearance of large assemblies of improperly annealed, partially double-stranded, heteroduplex DNA migrating above the library (known as 'daisy chains', Fig. 2B; see also Huppertz et al., 2014).
Adhere to the necessary safety measures when working with ethidium bromide.

Preparative PCR
From your cycle optimization PCR results, estimate the minimum number of PCR cycles to use to amplify the library. Consider that the template for the reaction will be 2.5-times more concentrated (see PCR mix below), therefore one cycle less is needed than in the scouting PCR (steps 105 to 108: PCR cycle optimization). 110. Run the same PCR program as in step 106, but with the adjusted cycle number.

Purification of the sequencing library
Prior to sequencing, all primers and amplicons that do not contain an insert need to be removed. Empty amplicons have a length of 140 bp, while amplicons with ribosome-protected fragment inserts exhibit a length of ∼170 bp. Here we provide instructions for library purification using an automated agarose gel electrophoresis system. If this is not available in the lab, the library can be purified via a native 8% polyacrylamide gel as described previously (McGlincy & Ingolia, 2017). 111. Purify the PCR reaction using the automated agarose gel electrophoresis system (PippinPrep, or similar) using a 3% agarose cassette according to the manufacturer's instructions with the following settings: target fragment length 168 bp with the setting 'tight'.

Quantitative and qualitative analysis of the purified sequencing library
For quality control and determination of the concentration of the libraries, a screen tape assay using the High Sensitivity D1000 ScreenTape and the High Sensitivity D1000 Reagents is employed on a TapeStation system.
113. Determine library concentration by measuring UV absorbance at 260 nm using the Nanodrop System.
114. Analyze an appropriate amount of the purified library using a high sensitivity screen tape assay on a TapeStation according to the manufacturer's instructions ( Fig. 2C and D).
For a standard experiment, we recommend to sequence at least 20 million reads per library of which typically 15 ± 5% map uniquely to the genome.
117. Extract unique molecular identifiers (UMIs) and append them to the read name using UMI-tools (version 1.0.1, extract-method=regex).
118. Filter reads by lengths and keep reads between 16 and 40 nt.
122. Analyze and interpret ribosome profiling data in R (version 3.5.1) and Rstudio using the riboWaltz package (version 1.2.0).

RIBOSOME PROFILING PROTOCOL FOR DIATOMS USING SUCROSE GRADIENT FRACTIONATION
This protocol includes the same steps as the Basic Protocol, but with the addition of a sucrose density gradient instead of a sucrose cushion. This allows users to optimize nuclease digestion conditions to suit a wider range of diatom species as monitored by the collapse of polysomes into monosomes after nucleolytic digestion.

Preparation of sucrose gradient
We recommend using a gradient master device (Biocomp Instruments) for rapid and reproducible formation of sucrose gradients. Here, 12 ml gradients are used with a SW41 rotor (Beckman). Conditions for monosome purification using different rotors are described in Meindl et al. (2023).
2. Pre-cool the ultracentrifuge and cool down the rotor and the buckets. Thaw all necessary reagents.
It is recommended to prepare the sucrose solutions well in advance to allow air bubbles to dissipate.
4. Prepare linear sucrose density gradients using the Biocomp gradient master according to the manufacturer's instructions. Use the 'long caps' provided by the manufacturer with the following program: SW41 rotor, sucrose, long caps, 10%-50%, 11 steps. If required, the density gradients can be stored at 4°C for several hours.
Sucrose gradient centrifugation 5. Remove the caps from the gradient tubes and gently overlay the solution with the sample. Pool replicates of the same sample at this point. Balance the tubes carefully with PRB buffer.
6. Carefully transfer tubes into the buckets, close lids and insert buckets into the rotor.

Density gradient fractionation
This protocol uses a gradient fractionation system which employs a tube piercing unit to deliver a dense chase solution to displace the gradient. It is connected to an FPLC system for analysis and fractionation. Further details are described in Meindl et al. (2023).
8. Connect the siFractor (siTOOLs Biotech) to an FPLC machine according to the manufacturer's instructions and thoroughly rinse the system with H 2 O. Fill a super loop (or similar) with sucrose chase solution and connect it to the FPLC system. Prime the tubes with the chase solution.
9. Follow the manufacturer's instructions for the fractionation of the samples from the gradient tubes while continuously monitoring conductivity and UV absorbance at 260 nm. Collect fractions of 1 ml each. Figure 2E depicts a typical UV profile obtained with nuclease treated T. pseudonana cell extract.
Pause point. Gradient fractions can be stored at −80°C for extended periods of time.

RNA isolation from density gradient fractions
10. Select the gradient fractions that contain your complexes of interest (e.g., 80S monosomes) and dilute 1:1 (v/v) with nuclease-free H 2 O to prevent phase inversion during organic extraction (due to the high sugar concentration of the sample).
11. For organic extraction, transfer the samples to a fume hood and add an equal volume of phenol/chloroform/isoamyl alcohol (25:24:1) and mix thoroughly by vortexing.
12. Separate the phases by centrifugation in a bench-top centrifuge for 10-30 min, 14,000 × g, room temperature, then transfer the upper aqueous phase into new tube. Make sure not to transfer any of the organic solution.
IMPORTANT: make sure not to transfer any of the interphase.
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14. Aspirate supernatant and wash the pellet with cold 80% ethanol. Aspirate the ethanol, air-dry the pellet and resuspend the RNA in 5 μl 10 mM Tris, pH 8.
Samples can be stored overnight at −20˚C or for several months at −80°C.
15. Continue with footprint fragment purification as outlined in the Basic Protocol step 18.

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Background Information
Ribosome profiling provides comprehensive snapshots of cellular translation based on the sequencing of mRNA fragments that are protected by translating ribosomes from nucleolytic digestion (Ingolia et al., 2009). These ribosome-protected fragments (RPFs) yield positional information of ribosomes on mRNA and their quantification allows the determination of protein synthesis rates. Moreover, by normalization to RNA abundance, ribosome loading scores can be derived that reflect the translation efficiency of mRNA species. Ribosome profiling monitors the final step of gene expression and therefore allows detection of gene expression changes that occur at different levels (including transcriptional regulation) (Ingolia, 2016). Changes in protein synthesis rates (as measured by changes in RPFs from a locus) can hence be driven, e.g., by transcriptional regulation, RNA turnover, differential RNA processing, or regulated translation. This allows to detect changes in gene expression programs, e.g., induced by a changing environment.
A wide range of ribosome profiling protocols are available for different species. Due to the different nature of cells and ribosomes, these protocols vary in their methods of harvesting, lysis conditions, and the selected nuclease and digestion conditions. In eukaryotic organisms, ribonuclease (RNase) I, A, T1 and micrococcal S7 are commonly used but their cutting efficiencies are species-dependent (Gerashchenko & Gladyshev, 2017). In this protocol, we used RNase I which performed well for T. pseudonana (Fig.  2E).
Different methods have been used in ribosome profiling studies to isolate digested monosomes, the most prevalent ones being ultracentrifugation via sucrose gradient and sucrose cushion (e.g., Ingolia et al., 2012;Meindl et al., 2023). As described in the Alternate Protocol, density gradient centrifugation was performed to identify optimal nuclease digestion conditions for T. pseudonana which had not been established before. Moreover, density gradient centrifugation allows the purification of ribosomal monosomes without carrying over a large portion of messenger ribonucleoproteins (mRNPs) to the next steps in the protocol (McGlincy & Ingolia, 2017). However, it requires additional instrumentation that may not be readily available in all labs.
Ribosome profiling sequencing libraries are typically dominated by rRNA fragments. Despite measures to limit these contaminations, e.g., precise excision of RNA in the right size after denaturing poly-acrylamide gel electrophoresis (PAGE) using appropriate markers, contaminations typically represent ∼90% of the reads. Hence, different measures have been implemented for their depletion. To specifically target the contaminants, they are experimentally identified typically by shallow sequencing of ribosome profiling libraries prepared under the same conditions from the respective organism/sample. Several commercially available rRNA depletion kits have been tested in a recent study and were found generally unsuitable for use with ribosome footprint libraries (Zinshteyn et al., 2020), especially kits based on targeted nuclease cleavage can lead to ribosome footprint degradation, reduction of mappable reads, interference with global gene expression measurements and blurred nucleotide resolution. Moreover, commercial kits that target T. pseudonana rRNA contaminations is not available. Thus, for our protocol we designed biotinylated antisense oligonucleotides that target the common contaminants found in sequencing libraries generated from ribosomal profiling of T. pseudonana and implemented a subtractive hybridization step in the protocol.
Various protocols for library preparation from ribosome profiling samples have been described that differ, e.g., in aspects of linker sequence and ligation, or the purification of the reaction intermediates. More recently, singlepot reactions have been introduced that exploit the template switching activity of reverse transcriptase (Ferguson et al., 2023;Ozadam et al., 2023). Here, we employ a recently developed protocol for sequencing library preparation that is particularly suited for low input samples (Meindl et al., 2023) and hence for the processing of samples derived from T. pseudonana that can have a low yield. In brief, for sequencing library preparation, an adapter sequence is ligated to the ribosome protected fragments, followed by reverse transcription, circularization of the cDNA and PCR amplification. Purification of the reaction intermediates occurs via solid phase reversible immobilization. Degenerate nucleotides on the ends of the adapters used in the ligation and circularization reactions reduce ligation bias and are employed for the identification of PCR duplicates during bioinformatic analyses. Bioinformatic analysis of ribosome profiling data has been described in detail elsewhere (e.g., McGlincy & Ingolia, 2017). We mapped the sequencing reads against the T. pseudonana genome instead of the transcriptome due to its higher quality.
Taken together, the adjustments made to previously existing protocols result in highquality sequencing data of actively translating ribosomes from T. pseudonana. Our protocol is robust and can now be used for future gene expression analysis in this organism, as well as a starting point for the adaption of the protocol to other related species.

Critical Parameters
For robust and reproducible results from ribosome profiling experiments several parameters are important, e.g., the way cells are harvested and lysed can have a profound effect on ribosome distribution by triggering of cellular stress responses (e.g., . Also, the use of translation inhibitors has been debated since it can introduce artefacts in particular species (Gerashchenko & Gladyshev, 2014;Sharma et al., 2021). Furthermore, the nuclease(s) and conditions for the generation of ribosome protected fragments need to be carefully chosen. Nucleases that work well in one species, might result in a complete loss of the sample in others (Gerashchenko & Gladyshev, 2017). Hence, the conditions under which ribosome profiling is performed must be carefully chosen and adapted to the model system and research question pursued. Our protocol has been optimized for T. pseudonana and yields high-resolution footprints with decent frame information (see Fig. 3B).
For other species, conditions suitable for ribosome profiling need to be experimentally established.
In all cases, ribosome profiling experiments need to be performed under conditions that prevent nucleolytic degradation of the samples, since shortening of the RPFs during library preparation compromises data quality. This includes working at low temperatures whenever possible, as well as the use of nuclease-free reagents and RNase inhibitors. Moreover, since the amount of input material for sequencing library preparation is typically rather limited, the use of low bind reaction tubes and low retention tips is highly recommended, once the RPFs have been excised from the gel. Experimental precision during critical steps, e.g., the size selection of the RPFs by denaturing page, limits contaminations and yields high quality data.
Proper amplification of the sequencing library is crucial for obtaining good results, since over-amplification increases the fraction of PCR duplicates in the final library. Yet, insufficient amplification of the sequencing library will produce only minute amounts of material that are often not sufficient for sequencing, the protocol provides a simple and robust step (scouting PCR) to identify optimal PCR conditions for the amplification of the library.

Troubleshooting
Since ribosome profiling is a complex technique with numerous factors contributing to success, troubleshooting of the experiments is complex. In the following, we provide a list of frequently encountered problems and how to deal with them.
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