Executing cell-specific cross-linking immunoprecipitation and sequencing (seCLIP) in C. elegans

Summary The single-end enhanced cross-linking immunoprecipitation (seCLIP) method is well suited for efficient and unbiased transcriptome-wide interrogation of RNA-binding protein (RBP) interaction sites. Here, we provide a protocol for executing cell-specific seCLIP for any desired RBP in Caenorhabditis elegans. We begin with steps and recommendations for transgene construction and Cas9-mediated chromosomal integration. We provide detailed procedures for isolation of RBP-associated RNA fragments, subsequent library preparation, and sequencing. We further discuss best practices for data analysis, interpretation of results, and troubleshooting. For complete details on the use and execution of this protocol, please refer to Blazie et al. (2021).1


SUMMARY
The single-end enhanced cross-linking immunoprecipitation (seCLIP) method is well suited for efficient and unbiased transcriptome-wide interrogation of RNA-binding protein (RBP) interaction sites. Here, we provide a protocol for executing cell-specific seCLIP for any desired RBP in Caenorhabditis elegans. We begin with steps and recommendations for transgene construction and Cas9-mediated chromosomal integration. We provide detailed procedures for isolation of RBP-associated RNA fragments, subsequent library preparation, and sequencing. We further discuss best practices for data analysis, interpretation of results, and troubleshooting. For complete details on the use and execution of this protocol, please refer to Blazie et al. (2021). 1

BEFORE YOU BEGIN
The goal of this method is to identify transcriptome-wide interaction sites for an RBP in specific cells in C. elegans. This method relies on expressing an epitope-tagged RBP transgene under a cell-specific promoter. In the following section, we outline recommended steps and considerations for designing RBP transgene expressing strains in preparation for seCLIP. seCLIP controls RBP binding sites are identified from seCLIP sequencing data as clusters of aligned sequencing reads. 2 However, many read clusters derive from non-specific association with common RNA 'contaminants' during immunoprecipitation, especially short, abundant transcripts (e.g., trans-splice leader transcripts and small nucleolar RNAs). Therefore, it is important to implement controls to help distinguish signal from noise in seCLIP sequencing data. At minimum, we recommend simultaneously performing seCLIP with a negative control strain lacking any RBP transgene (e.g., N2 strain), which is useful for identifying non-specific transcripts that stick to beads during immunoprecipitation (IP) (Blazie et al. 1 ). Sequenced read clusters identified from this 'no transgene' control can be regarded as background and subtracted from the RBP transgene data sets. We also recommend including a control transgene that expresses a mutated or truncated RBP lacking RNA binding activity. This 'inactive RBP' control is especially useful when the experimenter desires to pinpoint RNA binding sites for a specific RBP (target RBP) that co-immunoprecipitates with other RBPs (off-target RBPs). Performing seCLIP with the 'inactive RBP' control is crucial for identifying the off-target RBP-binding sites, which can be filtered from sample datasets to refine a list of high-confidence binding sites of the target RBP (see data analysis for more details).
Cas9 (derived from pDD162, 10 ), which facilitate CRISPR germline insertion of the transgene in the germline ( Figure 1B). Fluorescent markers expressed in the pharynx (Pmyo-2::mCherry) and muscle (Pmyo-3::mCherry) are also co-microinjected and serve to 1) identify successful microinjection events indicated by fluorescent F1 progeny, and 2) distinguish extrachromosomal array transgenics from single-copy insertion animals. The F1 progeny of the injected P0 animals are then treated with hygromycin and in several days single-copy insertion animals are identified on the basis of 1) resistance to hygromycin, and 2) lacking expression of the co-injection markers. Correctly integrated single-copy insertions can then be verified using three primers in a single PCR reaction. Two of the genotyping primers bridge the chromosomal insertion site (e.g., YJ10507/YJ10508 for Chr I) and yield a product from wild type alleles where no transgene insertion has ocurred. The third primer (YJ10686) anneals within the inserted hygromycin cassette and works with the chromosome specific primer (e.g., YJ10507 for Chr I) to yield a product from insertion alleles. Importantly, including all three primers in the PCR reaction will produce two different size products if both wild type and insertion alleles are present, allowing the user to distinguish between heterozygous and homozygous insertion animals.
In the following section, we describe the steps needed to generate a CasSCI transgenic line.
1. Clone the cell-specific transgene encoding epitope-tagged RBP into CasSCI vector for singlecopy genomic insertion. Note: We use Gibson Assembly technology 11 to clone the RBP transgene into the CasSCI vector. The choice of CasSCI vector (available in AddGene) will depend on the intended insertion site: use pCZGY2727 (for insertions on Chromosome I; ttTi4348) or pCZGY2729 (Chr IV; cxTi10882). Transgene cloning is facilitated by using SpeI and SphI restriction enzyme sites present on each CasSCI vector. See troubleshooting, problem 1 for advice on purifying the CasSCI vector.
a. Design oligonucleotides for transgene assembly (see Note below).
CRITICAL: Aim for at least 20 nucleotides and annealing temperature of $60 C for the 3 0 region of the primer annealing to the target sequence.
Note: We typically PCR amplify three DNA fragments corresponding to the promoter sequence, the epitope tag, and the RBP cDNA (including 3 0 UTR) with a high-fidelity (e.g., Phusion) polymerase. Design primers with 3 0 ends annealing to the target sequence. The 5 0 end of the primers should contain 20 additional nucleotides, which do not anneal to the target, but are instead homologous to the adjacent DNA fragment to facilitate Gibson assembly. For example, the 5 0 end of the forward primer used to amplify the promoter will contain 20 nucleotides homologous to the CasSCI vector sequence and the promoter reverse primer will contain 20 nucleotides homologous to the epitope tag-RBP. NEBbuilder is a useful tool for designing Gibson Assembly primers and checking plasmids designed with Gibson Assembly.
b. Digest the CasSCI vector (e.g., pCZGY2727) with SpeI and SphI (37 C for 1 h, then deactivate at 80 C for 20 min). c. Check products on a 1% standard agarose gel to ensure correct digestion (2 bands: $10.5 kb vector backbone and 1.8 kb insert). d. Prepare the Gibson assembly reaction (10 mL total volume): i. Add 1 mL each PCR product and 1 mL digested CasSCI vector to a PCR tube, then add ddH 2 O to 5 mL. ii. Add 5 mL Gibson Assembly Master Mix to the reaction. iii. Flick tube to mix, then briefly spin down.
Note: Gel extraction of the digested CasSCI vector backbone is not necessary as long as the SpeI and SphI enzymes were deactivated as instructed in step b. CRITICAL: We highly recommend setting up a control reaction with the Gibson assembly control DNA mix: 5 mL Gibson Assembly Control + 5 mL Gibson Assembly Master Mix. This can be used to verify the cloning reaction is working.
e. Incubate the reaction(s) at 50 C for 1 h. f. Transform 1 mL of the reaction into DH5-alpha competent cells and spread on Ampicillin LB plates. g. Screen several (typically 4) clones to verify correct assembly using restriction digest.
CRITICAL: We further recommend sequencing the final clone to verify mutations were not introduced during PCR.
Note: The amount of transgene to inject depends on the toxicity level of the RBP gene and should be determined empirically by titrating transgene concentration in microinjections until evidence of transgenesis (co-injection marker expression) is observed. The presence of sick or arrested F1 transgenic larval animals is a sign of transgene toxicity. As a guideline, we routinely obtained single-copy eif-3.G transgene insertion lines when injecting the transgene at 5 ng/mL, but very few transgene insertions when injecting more than 10 ng/mL. c. Microinject the DNA mixture into $30-40 P0 young adult hermaphrodite N2 animals following standard procedure (Mello and Fire, 1995). d. $1 h after microinjection, move the P0 animals to fresh 10 mm NGM plates seeded with OP50 (1-2 animals per plate) and culture at 25 C for 3 days. 3. Selection of transgenic animals.
a. After the incubation period, observe the plates under a fluorescent dissection microscope for evidence that the microinjections were successful.
Note: For 1-2 P0 animals plated, a good injection event will typically contain 15-20 fluorescent F1s. This step is merely to check that microinjection has worked. However, we recommend treating all plates (whether or not they contain fluorescent animals) with hygromycin as they may contain successful insertion events.
b. Prepare hygromycin solution (300 mL for each plate) by diluting 60 mL of the hygromycin stock (50 mg/mL) into 240 mL ddH 2 O (for 60 mm NGM plates; adjust accordingly if using larger plates). c. Add 300 mL hygromycin solution to each NGM plate with fluorescent animals.
i. Gently swirl the solution around the plates so it covers the plate surface.
ii. Leave the lids off the plates to dry for $15 min at room temperature (20 C-23 C). d. Culture the plates at 20 C for 3 days. e. After the incubation period, pick animals that survived hygromycin treatment and do not express the mCherry markers to new NGM plates.
CRITICAL: Hygromycin resistant animals should appear healthy and move normally. Many hygromycin sensitive animals will survive but become sterile or remain as larvae or dauer without expressing mCherry -avoid picking these! The presence of embryos in the adults is a good sign that the animal is hygromycin resistant. Also, true hygromycin resistant animals will often be young (L3-L4) at this stage as they can arise in the F2 progeny of F1 treated animals. Refer to troubleshooting, problem 2 for advice.
f. After $2 days, when hygromycin resistant animals have produced progeny, genotype the animals using PCR using primers according Table 1. Ensure the line is homozygous. g. Outcross the single-copy insertion transgenic line(s) as desired. This section describes steps for culturing and harvesting transgenic C. elegans, subjecting them to crosslinking to covalently fix the RBP to RNA, and lysing the animals to release cell contents for subsequent RNase treatment ( Figure 2A). The number of animals to culture for each seCLIP experiment will depend on several factors, including the RBP transgene expression level, the number of cells expressing the transgene, and the efficiency of immunoprecipitation of the target RBP. As a guideline, we obtained sufficient yields to build high-quality seCLIP libraries with $200 mL pelleted C. elegans expressing 33FLAG::EIF-3.G transgene in the cholinergic motor neurons (driven by Punc-17B). 1 Users should adjust the volume of animals depending on the transgene expression variables.
1. Culture each C. elegans strain on 12 large NGM plates (150 mm) seeded with OP50 bacteria at the required temperature for 3-4 days. a. Pick $10 adult hermaphrodites each onto 4 small seeded nematode growth media (NGM) plates (10 cm) and culture 20 C for $3 days. b. Slice the small plates into quarters and transfer these NGM chunks to 12 large 30 cm NGM plates. c. Plates are ready to harvest when they become confluent with C. elegans, nearly exhausting the bacterial lawn (typically $3-4 days at 20 C).
CRITICAL: Before harvesting, briefly examine the animals under a low power dissection microscope to ensure they appear healthy. It is not recommended to harvest C. elegans from plates after which the bacterial lawn is exhausted, as many of the animals will begin to experience starvation that could influence RBP activity.
Note: If desired, it is also possible to culture worms from specific larval stages using a modified protocol (for example 12  a. Centrifuge at 523 3 g for 2 min to pellet animals. b. Remove the M9 supernatant and add M9 media to 5 mL. c. Repeat steps a and b one time. 6. Prepare animals for crosslinking: a. Using a glass pipette, transfer animals onto two large (150 mm) unseeded NGM plates. b. Allow the liquid to evaporate and the animals to spread in an even lawn across the plate surface ($5-10 min).
Note: If needed, animals may be distributed onto more than two large unseeded NGM plates.
7. Pre-cool a small tabletop centrifuge (see key resources table) to 4 C. 8. Insert the animals into the Spectrolinker XL-1000 UV irradiator and initialize crosslinking using an energy setting of 3 kJ/m 2 .
CRITICAL: The success of crosslinking can be superficially judged by viewing the C. elegans plates under a dissection microscope. Crosslinked animals will be largely immobilized as though they are frozen in place. Pause point: Worm pellets may be flash frozen in liquid nitrogen and stored at À80 C for up to 2 months.
12. Resuspend the animals in 4 mL C. elegans lysis buffer and immediately transfer the tube to ice. 13. Sonicate the crosslinked C. elegans suspension with seven pulses (10 s each, power setting 11) with 50 s rest on ice in between pulses.
CRITICAL: Ensure the samples remain chilled on ice during the entire sonication procedure to avoid overheating the proteins.
14. Clarify lysates: a. Clear the lysates by spinning at 5,242 3 g for 5 min in the small tabletop centrifuge precooled to 4 C. b. Carefully move the cleared lysate (4 mL), splitting the volume into four 2 mL Eppendorf tubes (1 mL lysate each). c. Discard the insoluble pellet.
Note: Splitting the lysate volume into four tubes serves to promote better mixing during the following RNase treatment steps.
Pause point: Lysates may be flash frozen in liquid nitrogen and stored at À80 C for up to 1 week, if desired. Samples may be thawed exactly once when ready to proceed. Avoid multiple freeze thaws.
RNase treatment of C. elegans lysates Timing: 25 min In this step, clarified lysates are treated with RNase to fragment the total RNA in order to generate RBP associated RNA fragments that will be immunoprecipitated in subsequent steps ( Figure 2B).
CRITICAL: In all proceeding steps, 'gently flick to mix' refers to manually flicking the reaction tube and firmly tapping the tube on the table surface to gather the liquid to the tube bottom. Keep in mind that excessive force (such as vortexing) can damage RNA and enzymes.
15. Add 2 mL Turbo DNase to each 1 mL lysate sample, gently flick to mix, and return to ice. 16. Dilute RNase I 1:25 in 13 ice-cold PBS. 17. Add 10 mL of the diluted RNase I to each lysate and gently flick to mix. 18. Incubate the lysates in Thermomixer at 37 C for 5 min. 19. Immediately transfer the samples to ice, then add 11 mL Murine RNase Inhibitor and pipette to mix. 20. Centrifuge at 15,000 3 g at 4 C for 15 min. 21. Combine the four 1 mL RNase treated lysates reactions into two tubes (2 mL each), which will here forward serve as biological/technical replicates.
Immediately proceed to step 22.

Immunoprecipitation of RBP-RNA complexes
Timing: $16 h In seCLIP, RBP binding sites are identified based on the enrichment of read clusters mapped from the immunoprecipitated RBP (referred to as CLIP) versus an input control (designated as Input).
The following steps will describe cDNA library preparation from both CLIP and Input ( Figure 2C). We recommend generating and sequencing at least two biological replicate seCLIP libraries for all samples and controls.
Note: The anti-FLAG beads are too viscous to aspirate with most pipette tips. We recommend pipetting the bead suspension with a blunted P200 pipette tip cut with a razor.  Immediately proceed to step 41.

Timing: $22 h
Western blotting is used to make the target RBP-RNA complex visible so that the RNA can be isolated for cDNA preparation. The seCLIP strategy uses two PAGE gels: the imaging gel is used for western blot detection of the protein to verify the target RBP was successfully immunoprecipitated, and the preparative gel is used for isolation of the RBP-RNA complex. The Input and CLIP samples will be run on both imaging and preparative gels. The RBP-RNA complexes from Input and CLIP will subsequently be isolated from the preparative membrane, the proteins are removed with Proteinase K, and the RNA purified in preparation for subsequent cDNA library generation ( Figure 2E). Note: It is helpful to load the input and CLIP for each sample side by side (with diluted ladder mix in between). The alternating diluted ladder helps to space apart samples in the preparative gel to prevent cross contaminating sampes when protein-RNA complexes are isolated in step 65.
50. Load the imaging gel: a. Load 13 mL protein ladder (un-diluted) in the first lane. b. Load 15 mL of each imaging gel sample (both CLIP and Input) and save the remaining volume at À80 C as a back-up.
Note: it is not necessary to load diluted ladder between samples in the imaging gel, as the protein-RNA complex will not be excised from the imaging gel.  61. Wash the membrane 33 with 5% milk in 13 TBST for 5 min each at room temperature (20 C-23 C). 62. Add the secondary antibody at the appropriate concentration (1:5000 for Amersham NA934 anti-Rabbit secondary) in 10 mL total volume 5% milk in 13 TBST and incubate at room temperature (20 C-23 C) for 1 h with gentle rocking. 63. Wash the membrane: a. Gently rock membrane with 13 TBST (no milk) for 10 min. b. Remove the 13 TBST and add fresh 13 TBST. c. Repeat steps a and b three more times. 64. Develop the membrane with ECL using manufacturers recommendations (and image using western blot film of choice. CRITICAL: Take note of the position of the tagged-RBP in the IP samples relative to the protein ladder as this will be used in subsequent steps (Refer to Figure 3A for an example). Refer to troubleshooting, problems 3 and 4 for advice.
65. Excise the RBP-RNA complex: a. Place the preparative membrane on a glass surface. b. Using the developed imaging gel as a guide, slice the IP sample from the tagged-RBP band including 75 kD above this band (Refer to Figure 3A). c. Carefully slice this membrane into $1 mm strips, transfer all strips into a single clean 1.5 mL Eppendorf tube and place the tube on ice. 66. Prepare proteinase K (PK) mix on ice (200 mL per sample): 160 mL PK buffer + 40 mL Proteinase K Solution. 67. Prepare Urea/PK buffer: a. Dissolve 420 mg Urea in 500 mL PK buffer. b. Add PK buffer to a final volume of 1 mL. 68. Submerge the membrane slices in 200 mL of the PK mix and incubate for 5 min in the Thermomixer (1,200 rpm at 37 C Pause point: Samples may be stored at À80 C until ready to proceed.

Timing: 3 h (for steps 82 to 109)
This step dephosphorylates RNA in the Input samples and then ligates the InvRil19 adapter to the 3 0 RNA ends ( Figure 2F). The InvRil19 adapter will be subsequently used as a priming site for RT-PCR. CRITICAL: Save the remaining 5 mL in À20 C as backup. a. Magnetically separate beads and remove all liquid using a fine tip. b. Air dry the beads for 5 min at room temperature (20 C-23 C). 108. Resuspend the beads in 10 mL H 2 O and incubate at room temperature (20 C-23 C) for 5 min. 109. Magnetically separate beads and transfer the 10 mL supernatant to a new tube (this is your RNA).
Immediately proceed to step 110.

Generation of CLIP and input cDNA
Timing: 75 min (for steps 110 to 126) cDNA will be generated with the InvAR17 primers, which anneals to the InvRil19 adapter ligated to CLIP and Input RNA samples ( Figure 2G). a. Add 1 mL 80% EtOH, pipette mix, and move the resuspension to a new tube. b. Incubate at room temperature (20 C-23 C) for 30 s. c. Magnetically separate beads and remove the supernatant. 124. Repeat washes in 1 mL 80% EtOH (step 123) two more times. 125. Magnetically separate samples, remove all liquid with a fine tip, and air dry for 5 min. 126. Resuspend beads in 5 mL Tris-HCl (5 mM, pH 7.5) and incubate for 5 min at room temperature (20 C-23 C).

For ALL samples (CLIP and
CRITICAL: Do not remove liquid from beads. Immediately proceed to step 127. This step will ligate the InvRand3Tr3 adapter to the 3 0 end of the newly generated cDNA ( Figure 2H). InvRand3Tr3 will later serve as an annealing site for the D503_forward primer used to amplify cDNA libraries for sequencing on the Illumina instrument. InvRand3Tr3 additionally contains a short, randomized sequence (Unique Molecular Identifier) that serves as a barcode to later distinguish cDNA molecules generated from unique RNA fragments from PCR duplicated cDNAs during the computational analysis of sequencing data. qPCR is used to estimate the quantity of template cDNA in order to determine the number of PCR cycles needed for PCR amplification of the cDNA library. Note that it is not necessary to perform the qPCR in replicates. If desired to confirm accuracy of qPCR results, users can perform a duplicate PCR using a 1:100 dilution of the sample cDNA, which should yield a C T $3 cycles above the 1:10 cDNA dilution. After qPCR, cDNA libraries are generated using primers containing indexes for Single-end sequencing on the Illumina HiSeq instruments ( Figure 2I). The PCR reactions are cleaned-up using magnetic MyOne Silane beads to prepare for subsequent cDNA library size-selection and purification. In this step, cDNA libaries will be electrophoretically seperated on a 3% agarose gel, the DNA sizeselected (175-300 bp) and gel excised, and purified in preparation for single-end sequencing on the Illumina platform ( Figure 2J) Note: this will depend on the size of the gel box. Longer running times result in better resolution but require the user to cut larger agarose slices to purify the library.

Briefly image the gel under UV light.
Note: You should see a smear between $50 bp to 800 bp in sample lanes (Refer to Figure 3B for an example). See troubleshooting, problem 5 for advice.
CRITICAL: Minimize the time gels are exposed to UV as it can damage the cDNA.
164. With a clean razor blade, excise the gel slice from 175-350 bp and place into a 15 mL conical tube. 165. Excise and elute the gel using the Qiagen MiniElute gel extraction kit according to the following: a. Weigh the gel slice to determine volumes needed for following steps. b. Add 63 volumes of Buffer QG to melt the gel (e.g., for 100 mg gel slice, add 600 mL QG). c. Weigh the gel slice. d. Melt the gel at room temperature (20 C-23 C; do not heat, do not vortex) and gently shake tube to facilitate gel melting. e. Add 13 volume of the original gel of 100% isopropanol and mix well (100 mg gel = 100 mL isopropanol). f. Capture DNA in MiniElute column: i. Load 750 mL into a MiniElute column.
ii. Spin at max speed for 1 min.
iii. Repeat loading and spinning the sample as necessary until the entire sample has spun through the column. g. Wash the column once with 500 mL Buffer QG. h. Wash the column with Buffer PE: i. Add 750 mL Buffer PE.
ii. Spin at max speed for 1 min.
iii. Discard the flowthrough and spin column again for 2 min at max speed. i. Transfer the column to a fresh 1.5 mL Eppendorf tube and let it air dry for 2 min. j. Carefully add 12.5 mL Buffer EB to the center of the column, incubate for 2 min at room temperature (20 C-23 C), and spin at max speed for 1 min. Note: After trimming adaptors and filtering reads, CLIPper will align the remaining high-quality reads to the C. elegans reference genome to generate alignment maps (BAM files). CLIPper will identify RBP binding sites from the sequence maps on the basis of read cluster enrichment in CLIP versus Input samples ( Figure 2K) and outputs these clusters in .BED format (see quantification and statistical analysis below).

EXPECTED OUTCOMES
The success of cell-specific seCLIP in C. elegans should be carefully monitored during protocol execution. On the imaging western blot, the immunoprecipitated RBP will generally yield a smear between the expected RBP molecular weight and 75 kD above ( Figure 3A). However, RBP signals in the Input samples appear faint because the RBP concentration is much less than in CLIP (Figure 3A). Successful generation of cDNA libraries are indicated by a DNA smear on the agarose gel between $150-300 bp and relative absence of DNA in CLIP samples from the 'no transgene' control ( Figure 3B).

QUANTIFICATION AND STATISTICAL ANALYSIS
CLIPper identifies RBP binding sites from the alignment maps as statistically enriched read clusters (peaks) in CLIP samples relative to the Input control samples. CLIPper will output a raw list of RBP peaks along with their genomic position interval, enrichment (log foldchange) in CLIP/Input, and statistical confidence (P-value) in BED file format. The parameters in the BED files may be used to custom set statistical thresholds to prioritize RBP binding sites. As described above, it is often useful to perform seCLIP with a control RBP transgene deficient in RNA-binding activity (i.e., mutated RNAbinding domain) and a control strain without the RBP transgene (e.g., N2 strain). The RBP peaks detected from the control datasets can be considered non-specific background and ignored in the sample data set. The remaining peaks can be regarded with higher confidence as true signal.

LIMITATIONS
The sensitivity and specificity of cell-specific seCLIP highly depends on the efficiency of RBP immunoprecipitation. Key factors limiting IP success include the number of cells expressing the RBP, RBP expression level, RBP size and solubility, and the performance of the antibody used for IP. Therefore, we recommend considering which of these factors may limit your application and optimizing IP before implementing seCLIP. Users should also beware that seCLIP will inevitably yield some offtarget or artifactual RBP binding sites even when rigorous controls were included. It is therefore prudent to validate the most interesting RBP-binding sites with secondary experimental approaches.

Potential solution
We have sometimes observed poor miniprep efficiency of the CasSCI vector (without any DNA inserts of desired transgene) and have identified two solutions. 1) The CasSCI vectors contain a ccdB cassette and DB3.1 bacteria containing these plasmids propagate slowly. Grow the liquid bacteria culture (3 mL in LB with Ampicillin and Chloramphenicol) for 48 h (instead of 16 h) before miniprep. 2) When eluting DNA from the Qiagen miniprep column, add 50 mL of elution buffer preheated to 70 C and let the column sit at room temperature (20 C-23 C) for 2 min before spinning.

Problem 2
No single-copy insertion transgenic lines are obtained (all transgenic animals contain the extrachromosomal array markers and/or are sensitive to hygromycin) during CasSCI (before you begin, CasSCI transgenesis, step 3e).

Potential solution
Reduce the concentration of the CasSCI vector (containing the tagged-RBP) in the injection mix. We have obtained lines using between 5-15 ng/mL CasSCI vector, depending on the toxicity associated with RBP overexpression.
Inject into wild type (N2) hermaphrodites as mutant backgrounds could reduce CasSCI efficiency.
Inject at least 50 P0 hermaphrodites as the CasSCI insertion efficiency may be low for some transgenes.
Consider reducing the transgene size, if possible. We have observed reduced CasSCI insertion efficiencies with transgenes >3,500 bp.

Problem 3
Tagged-RBP is not detected on the imaging western blot (step 64).

Potential solution
If proteins are not observed on a western blot, we recommend quantifying the protein concentration resulting from C. elegans lysis (after step 14) using approaches such as Bradford or BSA assays. We ll OPEN ACCESS routinely obtain protein concentrations between 20-40 mg/mL from whole C. elegans lysates. We have observed that a clarified C. elegans lysate (after pelleting the lysis) will be golden brown in color and have used this to superficially judge the quality of lysis before subjecting samples to seCLIP. Lysates with poor protein yields will have a color closely resembling the starting lysis buffer (clear in color; not yellow or brown).
Ensure that protease inhibitors are added to the lysis buffer.
Check a large volume of the input lysate on a western blot to ensure that your RBP is sufficiently expressed and detectable.
Optimize concentrations of the primary and secondary antibody used for western blotting. Also, consider using a more sensitive western blotting detection agent (e.g., Femto ECL) for low abundance proteins.
Include a positive control tagged protein (it doesn't need to be an RBP) that has worked in previous western blotting applications to verify the IP and western blot reagents are working.

Problem 4
Additional signals of unexpected size appear on the western blot (step 64).

Potential solution
If genomic DNA was used to clone the RBP transgene, additional bands on the westen blot might indicate that the protein of interest encodes multiple protein isoforms. Consider performing seCLIP with an RBP trasngene encoded from a cDNA of the desired RBP protein isoform.
Ensure that fresh protease inhibitors were added to they lysis buffer to avoid RBP degradation.
Increase the salt concentration (NaCl) of the IP wash buffer and/or the number of washes after IP.
Optimize the concentration of the primary and/or secondary antibody used for western blotting. Excessive antibody concentration or incubation periods can result in high background.

Problem 5
No DNA smear is detected in the agarose gel in step 163.

Potential solution
Ensure that fresh RNAse inhibitors were added at each step where indicated.
Verify that all enzymes used to prepare the cDNA library (i.e., AffinityScript RT polymerase, RNA ligase, etc.) are working. Many commercial manufacturers will include a positive control that can be used to check enzyme performance. We recommend always using fresh enzyme stocks.

RESOURCE AVAILABILITY
Lead contact Further information and requests for resources and reagents should be directed to and will be reasonably fulfilled by the lead contact, Yishi Jin (yijin@ucsd.edu).

Materials availability
All genetic constructs and C. elegans strains are available upon request to the lead contact.