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Tethered MNase Structure Probing as Versatile Technique for Analyzing RNPs Using Tagging Cassettes for Homologous Recombination in Saccharomyces cerevisiae

Part of the Methods in Molecular Biology book series (MIMB,volume 2533)

Abstract

Micrococcal nuclease (MNase) originating from Staphylococcus aureus is a calcium dependent ribo- and desoxyribonuclease which has endo- and exonucleolytic activity of low sequence preference. MNase is widely used to analyze nucleosome positions in chromatin by probing the enzyme’s DNA accessibility in limited digestion reactions. Probing reactions can be performed in a global way by addition of exogenous MNase , or locally by “chromatin endogenous cleavage ” (ChEC ) reactions using MNase fusion proteins . The latter approach has recently been adopted for the analysis of local RNA environments of MNase fusion proteins which are incorporated in vivo at specific sites of ribonucleoprotein (RNP ) complexes. In this case, ex vivo activation of MNase by addition of calcium leads to RNA cleavages in proximity to the tethered anchor protein thus providing information about the folding state of its RNA environment.

Here, we describe a set of plasmids that can be used as template for PCR-based MNase tagging of genes by homologous recombination in S. cerevisiae . The templates enable both N- and C-terminal tagging with MNase in combination with linker regions of different lengths and properties. In addition, an affinity tag is included in the recombination cassettes which can be used for purification of the particle of interest before or after induction of MNase cleavages in the surrounding RNA or DNA. A step-by-step protocol is provided for tagging of a gene of interest, followed by affinity purification of the resulting fusion protein together with associated RNA and subsequent induction of local MNase cleavages.

Key words

  • RNA
  • RNP
  • Ribosome
  • Structure probing
  • Enzymatic probing
  • Micrococcal nuclease
  • Fusion protein
  • Chromatin endogenous cleavage
  • ChEC
  • Saccharomyces cerevisiae

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1 Introduction

Numerous methods have been developed over the years to chemically or enzymatically probe the structure of chromatin and of ribonucleoprotein (RNP ) complexes, two major manifestations of nucleic acid–protein complexes in eukaryotic cells [1,2,3,4]. Depending on the respective agent or enzyme applied, the accessibility, flexibility, secondary structure, or tertiary fold of the respective nucleic acid components can be analyzed either in vitro or in vivo. Combined with a high throughput sequencing readout these analyses can be performed genome- or transcriptome wide, respectively.

One of the enzymes which are routinely used to characterize nucleosome positions and chromatin states is the micrococcal nuclease (MNase) which is secreted by the bacterium Staphylococcus aureus. MNase has endo- and exonucleolytic activity which is strictly dependent on the presence of calcium. It cleaves both DNA and RNA with some preference for single stranded and for A/T- and A/U-rich regions (reviewed in [5,6,7]. In a typical nucleosome mapping experiment exogenous MNase is added to chromatin to perform a limited digest. Subsequently, positions of DNA cleavages and regions which were less accessible to the enzyme are determined. Information on the exact positioning of nucleosomes on DNA can be deduced if the size of a protected fragment is close to 146 base pairs which are typically protected by a nucleosome core particle [8,9,10].

Laemmli and colleagues have introduced a variation of this approach which is termed “chromatin endogenous cleavage ” (ChEC ) [11]. Here, a DNA-binding protein of interest is expressed in S. cerevisiae in fusion with MNase which remains inactive in the cell due to low calcium concentrations. Increasing the calcium concentration after cell breakage activates the enzyme which cleaves then close by accessible DNA. The resulting cuts at specific genomic loci can be analyzed by Southern blotting, or by high throughput sequencing for obtaining a genome wide data set [11,12,13,14]. ChEC and related MNase tethering approaches [15, 16] provide a valuable alternative to techniques as CHIP-Seq or DAM-ID [17, 18] for mapping of DNA binding sites of specific proteins. In addition, occupancies of fusion proteins at selected genomic loci can be analyzed in a quantitative way [13].

Protein components of RNPs have been as well expressed in fusion with MNase in yeast . They are potentially assembled in vivo at their endogenous location in the respective RNPs . The activation of MNase after cell breakage by the addition of calcium leads then to specific RNA cleavages in proximity to the fusion proteins which can be mapped with nucleotide resolution either by local [19] or by transcriptome wide primer extension analyses [20]. Experiments using a ribosomal model substrate showed that the radius of the enzymatic probe can be customized through introduction of differently sized linkers between the RNP-protein and MNase [19]. Cleavages up to around 6 nm surface distance from the anchor protein could be observed with longer linkers while more proximal cuts were strongly favored when using a short linker. Thus, in analogy to the ChEC methodology, probing of the RNA in proximity of a MNase fusion protein can reveal if, and in which region this protein binds to cellular RNPs . Importantly, additional information about the RNP ’s folding state in the surrounding of the MNase fusion proteins can be obtained. In this regard, the general accessibility of the substrate for the enzyme’s active center and the preference of MNase for single stranded and A/U-rich regions seem to be major determinants of spatially restricted cleavage positions [19, 20]. Consequently, a low number of defined cleavages can be expected in RNP ’s with a compact fold and thereby low accessibility. A/U-rich loops of spatially flexible hairpins which are often hardly detectable in structural analyses by X-ray crystallography or single molecule cryo-electron microscopy , were observed to be preferred substrates of MNase tethered to RNPs [20].

To further facilitate the application of MNase fusion proteins for both chromatin and RNP research we established a set of vectors that can be used as templates for PCR based chromosomal N- or C-terminal MNase tagging of genes by homologous recombination in S. cerevisiae . The tagging cassettes include regions coding for a variety of protein linkers, the MNase itself, a 3x-FLAG affinity tag as well as a heterologous selection marker (see Tables 1, 2 and 3). The cassettes were designed in a modular way allowing for straightforward exchange of each of the individual functional elements. This can be used to replace for example the rather strong RPS28B promoter present in the N-terminal tagging cassettes with other promoters better reflecting the endogenous expression level of the protein to be tagged. Too high expression levels might cause an excess of free MNase fusion proteins not assembled in RNPs or chromatin . This in turn could favor the appearance of nontethered cleavages. The plasmids further code for a set of different linker regions separating the anchor protein from the MNase in the fusion protein . The resulting protein linker regions vary in length from 6 to 63 amino acids and contain elements of different predicted conformational flexibility [22]. Finally, the cassettes contain sequences coding for the strong 3x-FLAG affinity tag which can be used for detection of the fusion proteins by Western blotting. In addition, the tag enables efficient affinity purification of the fusion proteins before MNase activation. This can be useful to facilitate the downstream analyses of the resulting cleavages in RNPs .

Table 1 Plasmids containing tagging cassettes
Table 2 Amino acid sequences of linker modules referred to in Table 1
Table 3 Primer design for PCR based homologous recombination

In the following, we provide instruction for PCR based MNase tagging of a gene of interest in S. cerevisiae using the described plasmid set. We then outline a step-by-step protocol for the analysis of MNase tagged RNPs , including their purification using the built-in 3x-FLAG tag and the subsequent activation of the tethered MNase to induce local RNA cleavages . For the use of generated fusion protein expressing strains in chromatin analyses we refer to recently published protocols [13, 14, 16, 23].

2 Materials

2.1 Template Plasmids for PCR Based Amplification of Tagging Cassettes

  1. 1.

    Plasmids which can be used as templates for amplification of recombination cassettes by PCR are described in Tables 1, 2, and 3. These plasmids and the corresponding sequence information are available upon request. Tables 4 and 5 provide more detailed information on how these plasmids were constructed.

  2. 2.

    Synthetic oligonucleotides required for PCR-based amplification of recombination cassettes are to be designed as outlined below in the methods section. Several manufacturers offer oligonucleotide synthesis services in the required size range (45–80 nucleotides) with sufficient quality in terms of purity and error rate. As example, EXTREmers offered by Eurofins/Genomics were successfully used for the tagging procedure described below.

Table 4 Plasmid construction
Table 5 Oligonucleotides used for plasmid construction

2.2 Generation by PCR of Tagging Cassettes for Homologous Recombination

  1. 1.

    Proofreading DNA Polymerase for PCR reactions with buffers provided by the manufacturer.

  2. 2.

    dNTP solution mix containing each dNTP in 10 mM concentration (NEB), stored at −20 °C.

  3. 3.

    10 M lithium chloride.

  4. 4.

    >99.8% ethanol (p.a.).

2.3 Transformation of Competent Yeast Cells and Screening for Positive Clones

  1. 1.

    Material required for transformation of yeast cells and denaturing extraction of proteins as described in detail in [25].

  2. 2.

    Material for size separation of DNA by native agarose gel electrophoresis according to [26].

  3. 3.

    YPD-Hygromycin plates: 1% (w/v) yeast extract (Difco Laboratories), 2% (w/v) Bacto Peptone (Difco Laboratories), 2% (w/v) glucose, 2% (w/v) agar. The medium is autoclaved for 20 min and supplemented with 200 μg/mL hygromycin B (ThermoFisher Scientific) before casting the plates. A 200 mg/mL stock solution of hygromycin B can be sterilized by filtration through a filter with a pore size of 0.22 μm (Sarstedt) and stored at −20 °C.

  4. 4.

    SC-URA plates: 0.67% (w/v) YNB with nitrogen (Sunrise Science Products), 0.062% (w/v) CSM Ura-Leu-Trp powder (Sunrise Science Products), 2% (w/v) glucose, 2% (w/v) agar. The medium is autoclaved for 20 min and supplemented with 100 mg/L leucine (Sunrise Science Products) and 50 mg/L tryptophan (Sunrise Science Products) before casting the plates. Stock solutions (50x) of these amino acids can be sterilized by filtration through a filter with a pore size of 0.22 μm (Sarstedt) and stored at 4 °C.

  5. 5.

    YPD medium: 1% (w/v) yeast extract (Difco Laboratories), 2% (w/v) Bacto peptone (Difco Laboratories), 2% (w/v) glucose. The medium is autoclaved for 20 min.

  6. 6.

    Material for SDS polyacrylamide electrophoresis and subsequent transfer of proteins to a PVDF membrane by the Western blotting approach according to [26].

  7. 7.

    Immobilon-P PVDF membrane for Western blotting (Carl Roth).

  8. 8.

    Primary anti-Flag-Tag antibody: Rat monoclonal anti-DYKDDDDK antibody, clone L5, 2 mg/mL (Agilent Technologies).

  9. 9.

    Peroxidase coupled secondary antibodies: Peroxidase-AffinityPure goat anti-rat IgG + IgM (H + L) (Dianova).

  10. 10.

    Buffer PBS: 137 mM sodium chloride; 2.7 mM potassium chloride, 10 mM disodium hydrogen phosphate, 2 mM potassium dihydrogen phosphate.

  11. 11.

    Buffer PBS-T: PBS with 0.05% Tween 20.

  12. 12.

    Dry milk powder.

  13. 13.

    BM-Chemiluminescence blotting substrate (Sigma-Aldrich) and an appropriate imaging system for chemiluminescence reactions.

2.4 Yeast Culture and Preparation of Cellular Extracts

  1. 1.

    YPD medium: 1% (w/v) yeast extract (Difco Laboratories), 2% (w/v) Bacto peptone (Difco Laboratories), 2% glucose. The medium is autoclaved for 20 min.

  2. 2.

    Protease inhibitor stock solution (100x concentrated): 200 mM benzamidine and 100 mM PMSF in ethanol p.a., stored at −20 °C.

  3. 3.

    Buffer AG100: 20 mM Tris–HCl pH 8.0, 100 mM potassium chloride, 5 mM magnesium acetate, 5 mM EGTA. Cool down the buffer to 4 °C and adjust it to 1x protease inhibitor concentration shortly before usage.

  4. 4.

    40 U/μL recombinant RNasin ribonuclease inhibitors (Promega), stored at −20 °C.

  5. 5.

    Glass beads with 0.75–1.0 mm diameter (Carl Roth).

  6. 6.

    Vibrax-VXR rotary shaker (IKA).

  7. 7.

    Protein assay dye reagent (Bio-Rad), stored at 4 °C.

  8. 8.

    Buffer AE+: 20 mM EDTA, 50 mM sodium acetate pH 5.3, stored at 4 °C.

2.5 Purification of Preribosomal Particles from Cellular Extracts and Induction of MNase Cleavage

  1. 1.

    Buffer AG100++: 20 mM Tris–HCl pH 8.09, 100 mM potassium chloride, 5 mM magnesium acetate, 5 mM EGTA, 0.1% (w/v) Tween, 0.5% (w/v) Triton X-100. Cool down the buffer to 4 °C and adjust it to 1× protease inhibitor concentration (see Subheading 2.4) shortly before usage.

  2. 2.

    Buffer AE+: see Subheading 2.4.

  3. 3.

    Poly-Prep chromatography columns with 10 mL reservoir (Bio-Rad).

  4. 4.

    Buffer AG100: see Subheading 2.4.

  5. 5.

    0.25 M calcium chloride.

  6. 6.

    1 mL spin columns with 35 μm filter (MoBiTec).

  7. 7.

    Water bath.

  8. 8.

    Turning wheel.

  9. 9.

    ANTI-FLAG ® M2 Affinity Gel (Sigma-Aldrich).

3 Methods

3.1 Primer Design and Choice of the Template Plasmid for PCR Reactions

Table 1 provides an overview on different plasmids which were created as templates for PCR based amplification of different tagging cassettes . Some of the plasmids are designed for N-terminal and others for C-terminal MNase tagging of a gene of interest in S. cerevisiae . Otherwise, the MNase cassettes encoded by the different plasmids differ by the linkers between MNase and the target protein and by the yeast selection markers. Two oligonucleotides have to be designed after choosing one of the plasmids as template for PCR based amplification of a recombination cassette (see for an overview [25, 27, 28]). These two oligonucleotides, in the following referred to as upstream and downstream oligonucleotide, are designed to contain at their 3′-end vector specific priming sequences indicated in the respective columns in Table 3. The priming sequences hybridize with corresponding regions on the plasmid which flank the tagging cassette and thus serve to amplify these cassettes in the PCR reaction. Sequence stretches of the gene of interest are added at the 5′-end of the primers. They should target cellular homologous recombination reactions of the amplified cassette to the gene of interest. For N-terminal tagging more than 40 nucleotides upstream of the start codon of the gene of interest are added 5′ to the upstream priming sequence to generate the upstream oligonucleotide. The start codon of the gene and more than 40 following nucleotides are added in reverse complement orientation to the 5′-end of the downstream priming sequence. Thus, the priming sequence in this downstream oligonucleotide directly follows 3′ after the reverse complement sequence of the start codon (5´-CAT-3′). For C-terminal tagging a stretch of more than 40 nucleotides ending with the last codon before the stop codon of the gene of interest is added 5′ to the upstream priming sequence in the upstream oligonucleotide. For the downstream oligonucleotide, more than 40 nucleotides of sequence just 3′ of the genes stop codon are added 5′ of the priming sequence in reverse complement orientation. Thereby the priming sequence continues directly 3′ after this reverse complement stretch of the genes 3′ untranslated region.

3.2 Generation of PCR-Based Tagging Cassettes for Homologous Recombination

  1. 1.

    Approximately 25 ng of the selected plasmid is used as template in PCR reactions.

  2. 2.

    PCR reaction mixtures are prepared with a pair of upstream and downstream oligonucleotides and a proofreading DNA polymerase according to the manufacturer’s instructions (see Note 1 ). To obtain sufficient amounts of the desired PCR product two to six reactions can be performed in parallel, each with 50 μL reaction volume (see Note 2 ).

  3. 3.

    The quantity and size of the PCR products are determined by native DNA agarose gel electrophoresis using 5 μL of the PCR reaction mixture. The expected sizes for tagging cassettes amplified from the different plasmid templates are indicated in Table 1.3.

  4. 4.

    Identical PCR reactions performed in parallel are pooled (see step 1) and supplemented with 0.1 × volume of 10 M lithium chloride and 2.5 volumes of ice-cold ethanol.

  5. 5.

    The samples are incubated for at least 30 min at −20 °C and subsequently centrifuged at 14,000 × g at 4 °C for at least 20 min.

  6. 6.

    The supernatant is discarded, and the remaining DNA pellet is resolved in 20 μL of water. Native DNA agarose gel electrophoresis [26] using about 10% of the DNA solution can be performed to confirm successful precipitation and to quantify the amount of obtained DNA.

3.3 Transformation of Competent Yeast Cells and Screening for Positive Clones

  1. 1.

    Yeast strains used for transformation with tagging cassettes based on plasmids K2515 and K2628 should not carry a hygromycin resistance marker gene. Yeast strains used for transformation with tagging cassettes based on K2372, K2373, K2488, K2489, K2490, K2491, K2510, and K2511 should have an inactivated URA3 gene. Strains expressing only Flag-tagged bait proteins without MNase can be generated using plasmid K2373. The latter strains can be included in the downstream RNP probing analyses as a negative control testing for MNase independent cleavage events in the RNP of interest.

  2. 2.

    Yeast cells are made chemically competent for transformation as outlined in detail in [25].

  3. 3.

    For homologous recombination, 50 μL of competent yeast cell suspension are transformed with 10 μg of the PCR amplified tagging cassette (see Subheading 3.2). Yeast transformation is performed as described in [25].

  4. 4.

    Cells transformed with recombination cassettes based on plasmids K2515 and K2628 are plated on YPD-Hygromycin plates. For all other recombination cassettes, cells are plated on SC-URA plates. Before spreading on YPD-Hygromycin plates, the transformed cells should regenerate for at least 6 h in 10 mL of liquid YPD medium at 30 °C with shaking. Plates are incubated upside down at 30 °C until appearance of colonies (2–5 days depending on the strain background and plates used).

  5. 5.

    Several [4,5,6,7,8,9,10] colonies are individually streaked out in about 1 cm × 1 cm large patches on respective new selective plates and incubated for 16–24 h at 30 °C.

  6. 6.

    The resulting cellular material is used for denaturing protein extraction which is performed according to [25].

  7. 7.

    Proteins extracted from the different clones are then separated by size using SDS polyacrylamide gel electrophoresis followed by Western blotting to a PVDF membrane [26].

  8. 8.

    The membrane is incubated for 1 h at room temperature in 5% (w/v) dry milk powder in PBS.

  9. 9.

    Immunodetection of fusion proteins is performed using rat anti Flag-Tag-antibodies (1:1000 diluted in PBST containing 1% dry milk powder) as primary antibodies and peroxidase coupled goat anti-rat antibodies as secondary antibodies (1:5000 diluted in PBST containing 1% dry milk powder). The membrane is washed three times for 5 min in PBST after each antibody incubation step.

  10. 10.

    Fusion proteins are visualized on the membrane using the BM Chemiluminescence Blotting Substrate according to the manufacturer’s instructions.

  11. 11.

    Clones expressing MNase fusion proteins of the expected size can be inoculated in YPD medium starting from the yeast cells streaked out in step 5 (see Notes 3 and 4).

3.4 Yeast Cell Culture and Preparation of Cellular Extracts

  1. 1.

    Yeast strains expressing MNase fusion proteins are cultured in YPD medium at 30 °C to an OD600 of approximately 0.8–1.2. For probing of (pre)ribosomal particles we routinely use culture volumes of 1–4 L for the subsequent analyses. The culture volume should be optimized for other RNPs depending on their respective expression level.

  2. 2.

    Cells are pelleted for 8 min with 9000 × g at room temperature.

  3. 3.

    The supernatant is discarded and the cells are resuspended in 20 mL of ice-cold water per liter of cell culture.

  4. 4.

    The suspension is transferred to a 50 mL reaction tube and centrifuged for 3 min at 4200 × g at 4 °C.

  5. 5.

    The supernatant is discarded. The cell pellet can be frozen at this stage at −20 °C and stored until usage. In this case the cells are thawed on ice before continuing with the next step.

  6. 6.

    At this point all buffers should be ready and cooled down to 4 °C. All subsequent steps are performed at 4 °C, unless stated otherwise.

  7. 7.

    The cells are resuspended in 10 mL of buffer AG100 per liter of cell culture.

  8. 8.

    The cells are centrifuged for 3 min at 4200 × g and 4 °C and the supernatant is discarded.

  9. 9.

    The wet weight of the cellular pellet is determined.

  10. 10.

    For each gram of cells 1.5 mL of buffer AG100 supplemented with RNasin (0.04 U/μL) is added and the cells are resuspended.

  11. 11.

    The cell suspension is distributed in portions of 800 μL to precooled 2 mL reaction tubes loaded with 1.4 g of glass beads.

  12. 12.

    For mechanical disruption of the cells, the tubes are shaken for 7 min at 4 °C on an IKA Vibrax basic shaker at maximum speed. Afterward, the cell suspensions are chilled on ice for 3 min. The procedure is repeated at least three times (see Note 5 ).

  13. 13.

    The cell lysate is centrifuged for 5 min at 14,000 × g and 4 °C.

  14. 14.

    The supernatant is pooled in a new 1.5 mL reaction tube and centrifuged for 10 min at 14,000 × g and 4 °C.

  15. 15.

    The supernatant is transferred to a new 1.5 mL reaction tube.

  16. 16.

    The protein concentration of the cleared cell extract is determined with the Bio-Rad protein assay according to the manufacturer’s instructions.

  17. 17.

    Here, samples can be taken for downstream RNA analyses. For this, a volume of cellular extract containing 300 μg of protein (= input sample) is added to 500 μL of ice-cold buffer AE+ and stored at −20 °C until usage (see Note 6 ).

3.5 Purification of RNPs from Cellular Extracts and Activation of MNase

  1. 1.

    The cellular extract prepared in 3.4 is supplemented with 0.1% (w/v) Tween and 0.5% (w/v) Triton X-100 (see Note 7 ).

  2. 2.

    Prior to affinity purification , the Anti-Flag M2 affinity matrix is equilibrated with buffer AG100++. For preribosomal particles we usually use 200 μL of matrix suspension containing about 100 μL matrix for cellular extracts prepared from 1 L of yeast cell culture. The suspension is transferred to Poly-Prep chromatography columns, washed once with 10 mL water and four times with 3 mL of buffer AG100++.

  3. 3.

    The equilibrated affinity matrix is transferred to the tube containing the cellular extract (step 1) and incubated for 1 h at 4 °C on a turning wheel.

  4. 4.

    The suspension is transferred into a Poly-Prep chromatography column and washed twice with 2 mL of buffer AG100++ and once with 10 mL AG100++.

  5. 5.

    The affinity matrix is suspended in 1 mL AG100 and transferred into a 1.5 mL reaction tube (see Note 8 ).

  6. 6.

    The bead suspension is centrifuged for 2 min at 2000 × g and 4 °C. A part of the supernatant is carefully taken off such that the affinity matrix remains in a volume of 200 μL in the reaction tube.

  7. 7.

    In order to activate the MNase , calcium chloride is added to a final concentration of 8 mM.

  8. 8.

    The reaction tube is incubated with gentle shaking. The optimal temperature and incubation time for the RNP analyzed is to be determined in pilot experiments (see Note 9 ).

  9. 9.

    500 μL of ice-cold AE+ is added to stop the reaction. The sample can then be stored at −20 °C until subsequent RNA analyses.

  10. 10.

    RNA extraction and the analyses of MNase-dependent RNA cleavages by either Northern blotting , targeted primer extension reactions or random primer extension analyses followed by high-throughput sequencing can be performed on samples thawed on ice as described in [20].

4 Notes

  1. 1.

    For PCR reactions we use routinely Herculase II Fusion DNA Polymerase (Agilent) according to the manufacturer’s instructions with the thermocycler programmed to the following settings: Initial melt for 2 min at 95 °C followed by a 35 cycles of 20 s melting at 95 °C, 20 s annealing at 45 °C and elongation for 2 min at 72 °C. Lastly, a final elongation step is performed for 5 min at 72 °C.

  2. 2.

    The template plasmids are designed in a modular way such that respective stretches of homology with the genomic region of interest can be cloned 5′ and 3′ of the plasmids tagging cassettes . Usage of gene fragments of 100–500 nucleotides size, whose synthesis is offered by several companies, and either classical [26] or more recent [29, 30] molecular cloning strategies allow to easily integrate these larger complementary sequence stretches into the plasmids listed in Table 1. In this way, PCR based amplification of the cassette can be avoided. Instead, the newly generated plasmid is amplified in E. coli , purified, and the cassette together with the flanking homology stretches can be excised with the appropriate combination of restriction enzymes. Due to larger stretches of homology with the target gene region a clear increase in the number of positive transformants can be expected when using this approach.

  3. 3.

    At least two clones expressing fusion proteins of the expected size should be chosen for further analyses. Identical cleavage pattern among these biological replicates minimizes the risk that the probing results are due to unexpected rare genetic events during homologous recombination.

  4. 4.

    Integration at the expected genomic locus can be further confirmed by PCR analyses on genomic DNA of the selected clone, using one primer pairing in the tagging cassette and another one pairing in reverse orientation at the endogenous genomic locus. In case that in the downstream analyses no cleavage pattern can be observed, the primers should be designed such that the MNase coding sequence is amplified. The identity of the MNase coding sequence can then be confirmed by DNA sequencing.

  5. 5.

    The protein concentration in the crude cell extract can be measured using the Bio-Rad protein assay according to the manufacturer’s recommendations, to test the disruption efficiency. Usual protein concentrations at this step should yield >10 mg/mL. If the concentration is lower, the procedure should be repeated until no significant further increase in protein concentration is observed.

  6. 6.

    At this point it is possible to perform RNP probing experiments in cellular extracts without prior affinity purification of the RNP of interest. For this, the cellular extract is supplemented with calcium chloride to a final concentration of 8 mM. After incubation at the desired temperature and time periods (see Note 9 ) a volume of cellular extract containing 300 μg protein is transferred to a new tube containing 500 μL of ice-cold buffer AE+. These samples can be stored at −20 °C or directly used for RNA extraction and downstream analyses.

  7. 7.

    For better handling, prepare 10% (w/v) stocks of both Triton X-100 and Tween 20 and use them to adjust the concentrations in the cellular extract.

  8. 8.

    At this point the suspension can be split into two equal volumes which are distributed to two 1.5 mL reaction tubes. They are treated in parallel as described below except that for one of the two reaction tubes step 7 (addition of calcium chloride) is omitted. This reaction serves then as control to determine calcium and MNase independent cleavages in the RNP analyzed.

  9. 9.

    To determine the optimal temperature and time period for incubation in the presence of calcium several reactions can be performed in parallel for which these parameters are varied. For preribosomal particles with intermediate maturation state we use 15 min incubation at 16 °C. At optimal incubation conditions a substantial amount of target RNA (>20%) should be cleaved as determined by RNA extraction and northern blotting . Excessive cleavage conditions should be avoided since they may favor disintegration of the analyzed RNP and accumulation of nontethered cleavages.

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Acknowledgments

We thank all members of the chair of Biochemistry III for their support of this work. This work was financially supported by the grant SFB 960 from the “Deutsche Forschungsgemeinschaft” (DFG).

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Correspondence to Joachim Griesenbeck , Herbert Tschochner or Philipp Milkereit .

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Teubl, F., Schwank, K., Ohmayer, U., Griesenbeck, J., Tschochner, H., Milkereit, P. (2022). Tethered MNase Structure Probing as Versatile Technique for Analyzing RNPs Using Tagging Cassettes for Homologous Recombination in Saccharomyces cerevisiae . In: Entian, KD. (eds) Ribosome Biogenesis. Methods in Molecular Biology, vol 2533. Humana, New York, NY. https://doi.org/10.1007/978-1-0716-2501-9_8

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