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. Author manuscript; available in PMC: 2022 Jan 1.
Published in final edited form as: Methods Mol Biol. 2021;2252:151–173. doi: 10.1007/978-1-0716-1150-0_6

Genome-wide Analysis of Translation in Replicatively Aged Yeast

Hanna Barlit 1,1, Manish K Rai 1,1, Sara Shoushtari 1, Carine Beaupere 1, Vyacheslav M Labunskyy 1,*
PMCID: PMC8565997  NIHMSID: NIHMS1751377  PMID: 33765274

Abstract

Protein synthesis is an essential process that affects major cellular functions including growth, energy production, cell signaling, and enzymatic reactions. However, how it is impacted by aging and how the translation of specific proteins is changed during the aging process remains understudied. Although yeast is a widely used model for studying eukaryotic aging, analysis of age-related translational changes using ribosome profiling in this organism has been challenging due to the need for isolating large quantities of old cells. Here, we provide a detailed protocol for genome-wide analysis of protein synthesis using ribosome profiling in replicatively aged yeast. By combining genetic enrichment of old cells with the biotin affinity purification step, this method allows large-scale isolation of aged cells sufficient for generating ribosome profiling libraries. We also describe a strategy for normalization of samples using a spike-in with worm lysates that permits quantitative comparison of absolute translation levels between young and old cells.

Keywords: Ribo-Seq, Mother enrichment program, Protein translation, Aging, Saccharomyces cerevisiae, Spike-in, Yeast

1. Introduction

Many age-related human diseases are associated with the loss of protein homeostasis including type 2 diabetes, cancer, neurodegeneration, and cardiovascular disease [1–3]. Despite the fundamental importance of protein translation in the aging process, relatively few studies have investigated translational changes in protein synthesis accompanying aging. Yeast Saccharomyces cerevisiae is one of the most established model systems to study eukaryotic aging given its short lifespan and amenability to genetic manipulations [4,5]. However, quantitative analysis of protein synthesis in replicatively aged yeast has been difficult due to the challenges of purifying large quantities of old cells [6].

Ribosome profiling (Ribo-Seq) has recently emerged as a powerful tool to monitor protein translation at the genome-wide level [7]. This method is based on deep sequencing of ribosome protected ~28 nt mRNA fragments, which allows genome-wide quantification of translation at nucleotide resolution [8,9]. In addition to global analysis of the actively translated regions of the transcriptome, combining ribosome profiling with total mRNA abundance measurements by RNA-Seq can also be used to estimate the translation efficiency of each transcript and contribution of both transcriptional and translational regulation to changes in gene expression between experimental conditions [10,11]. Although Ribo-Seq has been used to study translational changes and mechanisms of translational regulation in different physiological states as well as in response to physiological stress conditions (including nutrient limitation, heat shock, oxidative stress, endoplasmic reticulum stress), the standardized protocol for analyzing translational changes with aging is currently lacking. Here we describe a detailed ribosome profiling protocol used in our lab for analyzing changes in protein translation in replicatively aged yeast cells (Fig. 1). To isolate large quantities of aged cells sufficient for generating ribosome profiling libraries, we utilize the Mother Enrichment Program (MEP) [12]. This method permits efficient isolation of the old mother cells by using an estradiol-inducible system that leads to cell cycle arrest specifically in the daughter cells. By combining the MEP with affinity purification of biotin-labeled mother cells [13], we are able to routinely isolate more than 1X108 cells, which provides sufficient material to perform Ribo-Seq. Importantly, we compare changes in protein synthesis in replicatively aged cells with genetically identical young cells that were labeled with biotin and cultured under the same conditions, allowing direct comparison between young and old cells.

Fig. 1.

Fig. 1

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Overview of the Ribo-Seq protocol for analysis of translation in replicatively aged yeast cells. (a) Yeast cells are biotinylated and are grown in the presence of estradiol for 2 hrs (YNG) or 30 hrs (OLD), which induces the Mother Enrichment Program (MEP) preventing division of daughter cells. (b) Biotinylated mother cells are then separated using magnetic cell sorting enabling isolation of large quantities of old cells sufficient for generating ribosome profiling libraries. The age of yeast cells that have been isolated using the MEP and magnetic sorting is determined by counting the number of “bud scars” stained with calcofluor dye. The population of OLD cells obtained after sorting on average contains ~11–13 more bud scars per cell compared to YNG cells. (c) An equal amount of the worm lysate spike-in control (1%) is added into each sample proportional to the number of A260 units in yeast lysates allowing normalization of translation changes in aged and young cells.

Previous studies have also shown that the bulk protein synthesis is significantly reduced during aging in a range of organisms and different cell types [14] as well as in replicatively aged yeast cells [15]. In conditions when the global translation is decreased, normalization of raw sequencing reads to account for differences in overall rates of protein synthesis is required for absolute quantification of translation changes between samples. To overcome this limitation, we use a spike-in with the lysate prepared from Caenorhabditis elegans. For this, the yeast cellular extracts used for mRNA and ribosome footprint isolation are spiked with equal amounts of the worm lysate proportional to the quantity of total RNA in each sample. Adding this important spike-in control from the evolutionarily distant species allows us to quantify differences in overall changes of protein translation in replicatively aged yeast cells. This strategy can be directly applied for normalization of ribosome profiling experiments in other eukaryotic species or different cellular states to enable accurate quantification of in vivo translation.

2. Materials

2.1. Preparation of worm lysate

  1. C. elegans N2 strain.

  2. E. coli OP50 strain.

  3. 60 mm Petri plates.

  4. Nematode Growth Medium (NGM): 1.7% (w/v) agar, 50 mM NaCl, 0.25% (w/v) peptone, 1 mM CaCl2, 5 μg/mL cholesterol, 25 mM KPO4, 1 mM MgSO4.

  5. M9 buffer: 22 mM KH2PO4, 42 mM Na2HPO4, 86 mM NaCl, 1 mM MgSO4.

  6. Lysis Buffer: 20 mM Tris-HCl pH 8.0, 140 mM KCl, 5 mM MgCl2, 100 μg/mL cycloheximide, 0.5 mM DTT, 1% Triton X-100.

  7. Liquid nitrogen.

  8. 50 mL conical tubes.

  9. Screw cap 2 mL tubes.

  10. Non-stick 1.5 mL RNase-free tubes.

  11. BioSpec cryomill.

  12. Chrome-steel beads, 3.2 mm.

  13. Stainless steel microvials, 1.8 mL.

  14. Spectrophotometer.

2.2. Isolation of replicatively aged yeast cells

  • 1

    Yeast MEP strain (UCC8773, MATa his3Δ1 leu2Δ0 ura3Δ0 lys2Δ0 hoΔ::Pscw11-cre-EDB78-NatMX loxP-CDC20-intron-loxP-HphMX loxP-UBC9-loxP-LEU2) [16].

  • 2

    YPD medium: 1% yeast extract, 2% peptone, 2% glucose.

  • 3

    100 mg/mL Nourseothricin (Nat): prepare 100 mg/mL stock in sterile water, store at −20°C.

  • 4

    50 mg/mL Hygromycin B.

  • 5

    Refrigerated centrifuge (with a rotor for 50 mL conical tubes).

  • 6

    Refrigerated microcentrifuge.

  • 7

    Phosphate buffer saline (PBS).

  • 8

    EZ-Link Sulfo-NHS-LC-Biotin.

  • 9

    0.1 M Glycine-PBS: prepare 0.1 M glycine solution in PBS from 2.5 M glycine stock just before use.

  • 10

    PBS+BE: 1 mg/mL BSA, 2 mM EDTA in PBS.

  • 11

    1 mM 17β-estradiol: prepare a 1 mM stock in ethanol, store at −20°C.

  • 12

    Dynabeads Biotin Binder.

  • 13

    DynaMag magnet.

  • 14

    Resuspension Buffer: 20 mM Tris-HCl pH 8.0, 140 mM KCl, 5 mM MgCl2.

  • 15

    Liquid nitrogen.

  • 16

    50 mL conical tubes.

  • 17

    15 mL conical tubes.

  • 18

    Screw cap 2 mL tubes.

  • 19

    Non-stick 1.5 mL RNase-free tubes.

  • 20

    Automated cell counter.

2.3. Counting bud scars

  1. 4% Formaldehyde in PBS.

  2. Calcofluor (fluorescent brightener 28).

  3. Fluorescent mounting medium.

  4. Glass slides with coverslips.

  5. Fluorescence microscope with DAPI emission filter.

2.4. Preparation of yeast lysate

  1. Lysis Buffer: 20 mM Tris-HCl pH 8.0, 140 mM KCl, 5 mM MgCl2, 100 μg/mL cycloheximide, 0.5 mM DTT, 1% Triton X-100.

  2. Liquid nitrogen.

  3. 50 mL conical tubes.

  4. Screw cap 2 mL tubes.

  5. Non-stick 1.5 mL RNase-free tubes.

  6. BioSpec cryomill.

  7. Chrome-steel beads, 3.2 mm.

  8. Stainless steel microvials, 1.8 mL.

  9. Spectrophotometer.

2.5. Footprint extraction

  1. Ultracentrifuge with SW-41 Ti rotor.

  2. Thinwall polyallomer tubes, 13.2 mL.

  3. Gradient Buffer: 20 mM Tris-HCl pH 8.0, 140 mM KCl, 5 mM MgCl2, 100 μg/mL cycloheximide, 0.5 mM DTT.

  4. 10% Sucrose: prepare 10 % sucrose solution in Gradient Buffer just before use.

  5. 50% Sucrose: prepare 50 % sucrose solution in Gradient Buffer just before use.

  6. Chase Solution: 20 mM Tris-HCl pH 8.0, 140 mM KCl, 5 mM MgCl2, 60% sucrose.

  7. BioComp Gradient Master.

  8. RNase I, 100 U/μL.

  9. Head-over-heels rotator.

  10. Gradient fractionation system including tube piercer stand.

  11. UV monitor.

  12. Syringe pump.

  13. 0.5 mL centrifugal filters (100 kDa MWCO).

  14. Release Buffer: 20 mM Tris-HCl pH 7.0, 2 mM EDTA, 40 U/mL Superase-In.

2.6. Footprint fragment purification

  1. 20% SDS.

  2. Acid-phenol:chloroform, pH 4.5 (with IAA, 125:24:1).

  3. Non-stick 1.5 mL RNase-free tubes.

  4. 3M NaOAc, pH 5.5.

  5. 10 mg/mL Glycogen.

  6. Ethanol.

2.7. Poly(A) mRNA Extraction

  1. 20% SDS.

  2. Acid-phenol:chloroform, pH 4.5 (with IAA, 125:24:1).

  3. Non-stick 1.5 mL RNase-free tubes.

  4. 3M NaOAc, pH 5.5.

  5. 10 mg/mL Glycogen.

  6. Ethanol.

  7. Dynabeads Oligo (dT)25.

  8. DynaMag magnet.

  9. Binding Buffer: 100 mM Tris-HCl pH 7.5, 500 mM LiCl, 10 mM EDTA, 1% LiDS, 5 mM DTT.

  10. Washing Buffer A: 10 mM Tris-HCl pH 7.5, 150 mM LiCl, 1 mM EDTA, 0.1% LiDS.

  11. Washing Buffer B: 10 mM Tris-HCl pH 7.5, 150 mM LiCl, 1 mM EDTA.

  12. 10 mM Tris-HCl pH 8.0.

  13. 10X RNA fragmentation buffer: 100 mM ZnCl2, 100 mM Tris-HCl pH 7.0.

  14. RNase-free water.

2.8. Dephosphorylation

  1. T4 polynucleotide kinase (PNK), 10,000 U/mL.

  2. Superase-In RNase inhibitor, 20 U/μL.

  3. 10X TBE buffer: 1 M Tris-HCl pH 8.3, 0.9 M boric acid, and 0.01 M EDTA.

  4. 2X TBE-urea sample buffer.

  5. 15% polyacrylamide TBE-urea gels

  6. SYBR Gold nucleic acid gel stain.

  7. Blue light transilluminator.

  8. RNase-free disposable pellet pestles.

  9. Non-stick 1.5 mL RNase-free tubes.

  10. RNA elution buffer: 20 mM Tris-HCl, pH 7.0, 2.0 mM EDTA.

  11. 3M NaOAc, pH 5.5.

  12. 10 mg/mL Glycogen.

  13. Ethanol.

2.9. 3’-Adapter ligation

  • 21

    T4 RNA ligase 2 truncated KQ, 200,000 U/mL.

  • 22

    100 ng/μL Preadenylated 3’-adapter /5rApp/AGATCGGAAGAGCACACGTCT/3ddC/.

  • 23

    5’ Deadenylase, 10 U/μL.

  • 24

    Rec J exonuclease, 10 U/μL.

2.10. Reverse Transcription

  1. Superscript III reverse transcriptase, 200 U/μL.

  2. 10 mM Deoxynucleotides solution mix (dNTPs).

  3. 8 μM Reverse transcription (RT) primer
    • 5’-pGATCGTCGGACTGTAGAACTCTGAACGTGTAGATCTCGGTGGTCGCCGTATCATT /iSP18/GTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT-3’.

  4. 2M NaOH.

  5. 2M HCl.

  6. RNase-free water.

  7. 3M NaOAc, pH 5.5.

  8. 10 mg/mL Glycogen.

  9. Ethanol.

  10. 10XTBE buffer: 1 M Tris-HCl pH 8.3, 0.9 M boric acid, and 0.01 M EDTA

  11. 2X TBE-urea sample buffer.

  12. 10% polyacrylamide TBE-urea gels.

  13. SYBR Gold nucleic acid gel stain.

  14. Blue light transilluminator.

  15. RNase-free disposable pellet pestles.

  16. Non-stick 1.5 mL RNase-free tubes.

  17. 20 mM Tris-HCl pH 7.0.

2.11. Circularization and PCR Library Amplification

  1. RNase-free water.

  2. CircLigase II single-stranded DNA (ssDNA) ligase, 100 U/μL.

  3. 20 μM Forward PCR primer
    • 5’-AATGATACGGCGACCACCGAGATCTACACGTTCAGAGTTCTACAGTCCGACG-3’
  4. 20 μM Indexed Reverse PCR Primers (the index specific for each primer is underlined)
    • Index primer 1
    • CAAGCAGAAGACGGCATACGAGATCGTGATGTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT
    • Index primer 2
    • CAAGCAGAAGACGGCATACGAGATACATCGGTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT
    • Index primer 3
    • CAAGCAGAAGACGGCATACGAGATGCCTAAGTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT
    • Index primer 4
    • CAAGCAGAAGACGGCATACGAGATTGGTCAGTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT
    • Index primer 5
    • CAAGCAGAAGACGGCATACGAGATCACTGTGTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT
    • Index primer 6
    • CAAGCAGAAGACGGCATACGAGATATTGGCGTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT
    • Index primer 7
    • CAAGCAGAAGACGGCATACGAGATGATCTGGTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT
    • Index primer 8
    • CAAGCAGAAGACGGCATACGAGATTCAAGTGTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT
    • Index primer 9
    • CAAGCAGAAGACGGCATACGAGATCTGATCGTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT
    • Index primer 10
    • CAAGCAGAAGACGGCATACGAGATAAGCTAGTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT
    • Index primer 11
    • CAAGCAGAAGACGGCATACGAGATGTAGCCGTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT
    • Index primer 12
    • CAAGCAGAAGACGGCATACGAGATTACAAGGTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT

  5. Phusion high-fidelity DNA polymerase, 2,000 units/mL.

  6. 10 mM dNTPs.

  7. 5X DNA loading dye.

  8. 10X TBE buffer: 1 M Tris-HCl pH 8.3, 0.9 M boric acid, and 0.01 M EDTA.

  9. Non-denaturing 8% polyacrylamide TBE gels.

  10. SYBR Gold nucleic acid gel stain.

  11. Blue light transilluminator.

  12. RNase-free disposable pellet pestles.

  13. Non-stick 1.5 mL RNase-free tubes.

  14. 10 bp DNA ladder,1 μg/μL.

3. Methods

3.1. Preparation of worm lysate

  1. Culture 2,000–4,000 synchronized worms on lawns of E. coli OP50 on each NGM agar plate (3–5 plates) at 20°C to young adulthood (24 h after L4).

  2. Harvest worms by washing the plates with 2 mL M9 buffer.

  3. Transfer the mixture into 1.5 mL tubes. Wait 10 min to allow young adult worms to settle down at the bottom of the tube. Remove the supernatant containing eggs and bacteria.

  4. Resuspend the worms by adding 1 mL M9 buffer to each tube. Wait 10 min to allow young adult worms to settle down at the bottom of the tube, remove the supernatant. Repeat the wash two more times.

  5. Transfer washed worms into new 1.5 mL tubes (~500 μL worm pellet/tube). Remove the supernatant and resuspend in 1 mL Lysis Buffer. Slowly dispense the mixture into the 50 mL tube with liquid nitrogen. Transfer the flash-frozen droplets into 2 mL screw-cap tubes and store them at −80°C.

  6. Pre-chill a 1.8 mL stainless steel tube with 3 chrome-steel beads in liquid nitrogen. Add 0.35–0.4 g frozen worm pellets, cover with a silicone rubber cap.

  7. Homogenize frozen worm pellets by cryogrinding for 10 sec at 4,200 rpm, immediately chill the tube in liquid nitrogen. Repeat 10 times.

  8. Add 1 mL of Lysis Buffer, mix well by pipetting. Transfer into a new 1.5 mL tube.

  9. Remove debris by centrifugation at 20,000 x g for 5 min at 4°C, transfer the supernatant into a new 1.5 mL tube.

  10. Determine OD260 and dilute the worm lysate to 50 OD260 units/mL with Lysis Buffer. Transfer 50 μL aliquots of the worm lysate into new tubes (2.5 OD260 units/tube). Flash freeze in liquid nitrogen, and store at −80°C. Prepare a large batch that can be used for multiple experiments.

3.2. Isolation of replicatively aged yeast cells

  1. Starting from a single colony, grow the MEP strain in 5 mL of YPD supplemented with 100 μg/mL Nat and 300 μg/mL hygromycin B (to maintain selection) at 30°C overnight.

  2. Measure the OD600 and dilute the overnight culture to OD600 ~ 0.2 in 40 ml YPD supplemented with 100 μg/mL Nat and 300 μg/mL hygromycin B. Divide the diluted culture equally into two 50 mL conical tubes.

  3. Culture cells at 30°C with shaking until OD600 reaches 0.6 (log phase).

  4. Harvest cells in 50 mL tubes in a refrigerated centrifuge at 3,000 x g, 4°C for 3 min.

  5. Wash cells 2 times in 10 mL of sterile PBS.

  6. Resuspend cells in 1 mL PBS.

  7. Dilute 1 μL of the cell suspension to 50 μL by adding 49 μL of H2O (50-fold dilution). Determine the cell density using an automated cell counter or a hemocytometer.

  8. Transfer 3×108 cells to a new 1.5 mL tube for labeling with biotin, centrifuge at 2,000 x g, 4°C for 3 min, and remove the supernatant.

  9. Warm-up EZ-link Biotin to room temperature before opening. Prepare 10 mg/mL EZ-link Biotin stock solution in PBS and add 300 μL to cells from the previous step (100 μL per 1×108 cells).

  10. Rotate the tube for 30 min at room temperature.

  11. Wash biotin-labeled cells two times in 1 mL 0.1 M Glycine-PBS to quench and remove free biotin.

  12. Centrifuge the sample at 2,000 rpm (400 x g), 4°C for 2 min (see Note 1).

  13. Resuspend the cells in 1 mL PBS and divide them into two 1.5 mL tubes (1.5 × 108 cells/tube). Keep on ice until inoculation.

  14. Inoculate 1.5×108 labeled cells into 20 mL of cold YPD media containing 1 μM 17β-estradiol and grow at 30°C for 2 h. These cells will be used to isolate the young cells sample (YNG) (Fig. 1, see Note 2).

  15. For isolation of old cells inoculate 1.5 × 108 labeled cells from step 13 to 700 mL of cold YPD media and add 17β-estradiol to 1 μM final concentration to induce the MEP. Grow cells at 30°C for 20 h, then add 300 mL of fresh YPD media containing 1 μM 17β-estradiol and continue growing at 30°C for an additional 10 h (30 h total incubation time). These cells will be used to isolate the old cells sample (OLD) (Fig. 1, see Note 3).

  16. After the incubation is completed, harvest cells by centrifugation in 50 mL tubes in a refrigerated centrifuge at 3,000 x g, 4°C for 3 min. Remove the supernatant.

  17. Wash cells twice in 30 mL of cold PBS. Resuspend washed cells in 1 mL (YNG) or 10 mL (OLD) of PBS+BE, and transfer to a 1.5 mL tube (YNG) or a 15 mL conical tube (OLD), respectively (see Note 4).

  18. Resuspend Dynabeads Biotin Binder before use. Transfer 300 μl of Dynabeads Biotin Binder (1.2×108 beads) into a 1.5 mL tube for YNG cells and 1 mL (4×108 beads) for OLD cells. Remove the buffer by placing on the magnet for 30 sec and discard the supernatant.

  19. Wash Dynabeads Biotin Binder in each tube 2 times in 1 ml PBS+BE buffer.

  20. Add an aliquot of Dynabeads Biotin Binder to YNG and OLD cells from step 16 and incubate at 4°C for 1 hr with rotation.

  21. Place the tube with YNG cells on the magnet for 30 sec, carefully remove the supernatant without disturbing the beads/labeled cells attached to the magnet. Proceed to washing the beads (step 22). For OLD cells, transfer a 1.5 mL aliquot of cell suspension into a 1.5 mL tube. Place the tube on the magnet for 2 min and slowly remove unlabeled cells without disturbing the beads/old cells attached to the magnet. Add another aliquot of cells to the tube. Repeat this process several times until all OLD cell suspension is processed.

  22. Wash the beads in each tube 3 times with 1 mL of cold PBS. Transfer the beads/labeled cells into an RNase-free 1.5 mL tube.

  23. Repeat step 22 two more times.

  24. Resuspend the cells in 1 mL YPD. Incubate the cells with shacking at 30°C for 30 min to recover.

  25. Dilute 1 μL of cells to 10 μL by adding 9 μL of H2O (10-fold dilution). Determine the cell density using an automated cell counter or a hemocytometer. Calculate the yield obtained after cell sorting. For both YNG and OLD samples, save 10 μl of cells for counting bud scars (see Note 5).

  26. Centrifuge cells 3,000 rpm (800 x g) 3 min. Remove the supernatant. Resuspend cells in 150 μL of Resuspension Buffer, slowly dispense the mixture into the 50 mL tube with liquid nitrogen. Transfer the flash-frozen droplets into 2 mL screw-cap tubes and store at −80°C.

3.3. Counting bud scars

  1. For counting bud scars, mix 10 μl of cells with 1 mL of 4% formaldehyde (in PBS), rotate for 10 min at room temperature.

  2. Centrifuge at 3,000 rpm (800 x g) 3 min and remove the supernatant.

  3. Wash cells twice with 1 mL PBS. Resuspend cells in 400 μL PBS containing 1 mg/mL calcofluor. Prepare 1 mg/mL calcofluor solution in PBS from 10 mg/mL stock just prior to use. Incubate cells for 15 min in the dark at room temperature.

  4. Wash cells once with 1 mL PBS. Centrifuge at 3,000 rpm (800 x g) 3 min and resuspend in 20 μL H2O.

  5. Mix 5 μL of cells with 5 μL of fluorescent mounting media, place on a glass slide and cover with a coverslip. Image the stained bud scars on a fluorescence microscope using a DAPI emission filter. If cell sorting is successful, OLD cells should contain on average ~11–13 more bud scars per cell compared to YNG cells (Fig. 1).

3.4. Preparation of yeast lysate

  1. Prepare several tubes containing flash-frozen droplets of Lysis Buffer (250 μL per tube) by slowly dispensing the buffer into the 50 mL tube with liquid nitrogen. Transfer the flash-frozen droplets into 2 mL screw-cap tubes and store them at −80°C until cryogrinding.

  2. Pre-chill a 1.8 mL stainless steel tube with 3 chrome-steel beads in liquid nitrogen. Add frozen yeast pellets (Section 3.2 step 26) (see Note 6).

  3. To the same tube add 250 μL of flash-frozen Lysis Buffer droplets, cover with a silicone rubber cap.

  4. Homogenize yeast cells by cryogrinding for 10 sec at 4200 rpm, immediately chill the tube in liquid nitrogen. Repeat 10 times.

  5. Add 500 μL of Lysis Buffer, mix well by pipetting. Transfer into an RNase-free 1.5 mL tube.

  6. Remove debris by centrifugation at 20,000 x g for 5 min at 4°C, transfer the supernatant into a new 1.5 mL tube.

  7. Determine OD260 of the yeast lysate. Calculate the amount of the worm lysate spike-in control (Section 3.1 step 10) required for each sample. Add 1 A260 unit of the worm lysate per 100 A260 units of each yeast lysate (to a final concentration 1%) for normalization (see Note 7).

  8. Divide the lysate equally into two aliquots that will be used for the preparation of “footprint” and “mRNA” libraries. Flash freeze in liquid nitrogen, and store at −80°C. The lysates can be stored at −80°C indefinitely.

3.5. Footprint extraction

  1. Pre-chill the SW-41 Ti rotor with buckets to 4°C.

  2. Prepare linear sucrose gradients (10–50% sucrose) in Gradient Buffer in 13.2 mL thin-wall polyallomer tubes using BioComp Gradient Master following the manufacturer’s instructions (see Note 8). Keep the gradients at 4°C until use.

  3. Thaw an aliquot of yeast lysate (“footprint” sample) on ice. Calculate the amount of RNase I (100 U/μL) needed for digestion for each sample. Add 20 U of RNase I per each A260 unit of yeast lysate and incubate 1 hr at room temperature with gentle rotation on a head-over-heels rotator.

  4. Centrifuge lysate at 20,000 x g for 5 min at 4°C to remove debris. Transfer the supernatant into an RNase-free 1.5 mL tube and put on ice.

  5. Load lysate on top of a 10–50% sucrose gradient.

  6. Centrifuge the gradients at 35,000 rpm (151,000 x g) at 4°C for 3 hrs using the ultracentrifuge with the SW-41 Ti rotor (see Note 9).

  7. Turn on the Gradient fractionation system and UV monitor. Set the range on UV monitor to 2.0 for maximum detection limit.

  8. Carefully transfer the tube with the sucrose gradient into the piercer stand. Fill the syringe with Chase Solution and connect it to the piercer stand tubing.

  9. Pierce the tube and start pumping Chase Solution at 1 mL/min using the syringe pump. By monitoring UV absorbance at 254 nm, collect fractions corresponding to the monosome peak (Fig. 2), and put on ice.

  10. Repeat fractionation for the rest of the samples (see Note 10).

  11. Concentrate the monosome fractions using 0.5 mL centrifugal filters (100 kDa MWCO). For this, load the monosome fractions into the centrifugal filters, centrifuge at 12,000 x g for 10 min at 4°C. Discard the flow-through, repeat until the volume reaches 100 μL.

  12. Add 400 μL of Release Buffer. Pipette up and down to mix and incubate 10 min on ice. Transfer the filter unit into an RNase-free 1.5 mL collection tube.

  13. Centrifuge at 12,000 x g for 10 min at 4°C. Do not discard the flow-through.

Fig. 2.

Fig. 2

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Sucrose gradient fractionation of the yeast lysates prepared from varying quantities of old cells. Cell lysates containing the indicated number of A260 units (a) or varying numbers of cells (b) were digested with RNase I and fractionated using 10–50% sucrose gradients. In our experience, we were able to collect the monosome peak and successfully prepare Ribo-Seq libraries from as low as 50×106 cells.

3.6. Footprint fragment purification

  1. Collect the flow-through (400 μL) containing footprint RNA fragments and transfer to an RNase-free 1.5 mL tube. Add 20 μL of 20% SDS (1% final concentration), pipette up and down to mix.

  2. Add 400 μL of acid-phenol:chloroform and vortex for 10 sec (see Note 11).

  3. Heat tubes at 65°C for 5 min, put on ice for 1 min. Centrifuge at 12,000 x g for 5 min at 4°C, transfer the aqueous layer (~350 μL) to an RNase-free 1.5 mL tube.

  4. Add 35 μL of 3 M NaOAc, 3.5 μL of glycogen, and 2.5 volumes of 100% ethanol. Incubate at −20°C for at least 1 hr to precipitate RNA.

3.7. Poly(A) mRNA Extraction and mRNA fragmentation

  1. Thaw an aliquot of cell lysate (“mRNA” sample) on ice, add 20 mM Tris-HCl pH 7.0 to bring the volume to 500 μL. Add 25 μL of 20% SDS (1% final concentration), pipette up and down to mix.

  2. Heat tubes at 65°C briefly to dissolve SDS. Add 1 volume of acid-phenol:chloroform, vortex for 10 sec (see Note 11).

  3. Heat tubes at 65°C for 5 min, put on ice for 1 min. Centrifuge at 12,000 x g for 5 min at 4°C, transfer the aqueous layer (~400 μL) to an RNase-free 1.5 mL tube.

  4. Repeat phenol extraction. Add 1 volume of acid-phenol:chloroform, vortex for 10 sec. Heat tubes at 65°C for 5 min, put on ice for 1 min. Centrifuge at 12,000 x g for 5 min at 4°C, transfer the aqueous layer (~350 μL) to an RNase-free 1.5 mL tube.

  5. Add 35 μL of 3 M NaOAc, 3.5 μL of glycogen, and 2.5 volumes of 100% ethanol. Incubate at −20°C for at least 1 hr to precipitate RNA.

  6. Centrifuge the samples at 20,000 x g for 30 min at 4°C to pellet the RNA, remove the supernatant. Centrifuge at 20,000 x g for 30 sec, remove the rest of the supernatant with a gel-loading tip, and air-dry the pellet for 5 min.

  7. Dissolve the pellet in 300 μL of Binding Buffer, pipette up and down to mix.

  8. Resuspend Dynabeads Oligo(dT)25 before use. Transfer 250 μL of beads to an RNase-free 1.5 mL tube and place the tube on a magnet. Wait 30 seconds and remove the supernatant.

  9. Take the tube out of the magnet and wash beads with 250 μL of fresh Binding Buffer. Remove Binding Buffer by placing the tube on the magnet, wait for 30 seconds and remove the supernatant.

  10. Add 300 μL of the RNA sample (step 7), mix by pipetting. Incubate for 5 min at room temperature with continuous mixing on a head-over-heels rotator.

  11. Place the tube on the magnet, wait for 2 min and remove the supernatant.

  12. Wash beads twice with 600 μL of Washing Buffer A at room temperature. Place the tube on the magnet to separate beads from the supernatant between each wash.

  13. Wash beads twice with 300 μL of Washing Buffer B at room temperature. Place the tube on the magnet to separate beads from the supernatant between each wash.

  14. Remove the washing buffer. Add 20 μL of 10 mM Tris-HCl pH 8.0 and incubate at 65°C for 2 minutes. Place the tube on the magnet, transfer the supernatant containing mRNA to a new 1.5 mL tube and put on ice. Do not discard the beads.

  15. Wash beads twice with 300 μL of Washing Buffer B.

  16. Dilute the mRNA with 4 volumes of Binding Buffer (e.g., if mRNA is eluted in 20 μL, add 80 μL of Binding Buffer).

  17. Place the tube on the magnet, wait for 2 min and remove the supernatant.

  18. Add the diluted mRNA and incubate with continuous mixing on a head-over-heels rotator for 5 min.

  19. Repeat steps 11–13.

  20. Remove the washing buffer. Add 20 μL of 10 mM Tris-HCl pH 8.0 and incubate at 65°C for 2 minutes. Immediately place the tube on the magnet, transfer the supernatant containing the mRNA to a new RNase-free tube.

  21. Add 20 μL of RNase-free water, 4 μL of 3M NaOAc, 1 μL of glycogen, and 2.5 volumes of 100% ethanol. Incubate at −20°C for at least 1 hr to precipitate RNA.

  22. Centrifuge the samples at 20,000 x g for 30 min at 4°C to pellet the RNA, remove the supernatant. Centrifuge at 20,000 x g for 30 sec, remove the rest of the supernatant with a gel-loading tip, and air-dry the pellet for 5 min.

  23. Resuspend mRNA in 18 μL of RNase-free water. Add 2 μL of 10X RNA fragmentation buffer, incubate at 94°C for 5 min, and put on ice.

  24. Add 20 μL of RNase-free water, 4 μL of 3M NaOAc, 1 μL of glycogen, and 2.5 volumes of 100% ethanol. Incubate at −20°C for at least 1 hr to precipitate RNA.

3.8. Dephosphorylation

  1. Centrifuge “footprint” and fragmented “mRNA” samples at 20,000 x g for 30 min at 4°C, remove the supernatant. Centrifuge at 20,000 x g for 30 sec, remove the rest of the supernatant with a gel-loading tip, and air-dry the pellet for 5 min.

  2. Resuspend the pellet in 7.75 μL of RNase-free water. Add 1 μL of 10X T4 PNK buffer, 1 μL of T4 PNK, and 0.25 μL of Superase-In. Pipette up and down to mix and incubate at 37°C for 1 hr.

  3. Rinse wells of the 15% TBE-urea gel from urea and pre-run the gel at 180 V for 15 min in 1X TBE buffer.

  4. Add 10 μL of 2X TBE-urea sample buffer to 10 μL of each “footprint” and “mRNA” sample.

  5. Prepare an oligo sizing control containing 1 μL of 10 μM 32nt RNA oligonucleotide, 1 μL of 10 μM 28nt RNA oligonucleotide, 8 μL water, and 10 μL of 2X TBE-urea sample buffer for “footprint” samples. Prepare an oligo sizing control containing 1 μL of 10 μM 63nt RNA oligonucleotide, 9 μL water, and 10 μL of 2X TBE-urea sample buffer for “mRNA” samples.

  6. Heat the samples at 75°C for 3 min, spin down at max speed for 10 sec, put on ice for 1 min. Load each sample into 2 wells of the 15% TBE-urea gel, run the gel at 180 V for 1 hr.

  7. Dissolve 5 μL of SYBR Gold in 50 mL of RNase-free water. Stain the gel with SYBR Gold for 5 min, protect from light.

  8. Using a blue light transilluminator, cut the gel slices between 28 and 32 nt markers for ‘footprint” samples and ~ 50–70 nt for “mRNA” samples with a razor blade (Fig. 3). Freeze the polyacrylamide gel slices at −80°C for 10 min.

  9. Heat the gel slices at 70°C for 2 min, grind the gel using RNase-free disposable pellet pestles. Add 300 μL of RNA elution buffer, 1 μL Superase-In, and incubate at 37°C for 3 hrs to extract RNA.

  10. Centrifuge the samples at 12,000 x g for 5 min at 4°C and transfer the supernatant to a new 1.5 mL tube. Add 30 μL of 3 M NaOAc,3 μL of glycogen, and 2.5 volumes of 100% ethanol. Incubate at −20°C for at least 1 hr to precipitate RNA.

Fig. 3.

Fig. 3

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Representative footprint and fragmented mRNA size selection gels after T4 polynucleotide kinase treatment. (a) Footprint fragments prepared from YNG and OLD yeast samples according to our protocol were separated on 15% polyacrylamide TBU-urea gel. 28 nt and 32 nt RNA oligonucleotides are used to guide the size of the gel slice that should be excised. (b) For fragmented mRNA samples, cut the gel slice around 50–70 nt. 63-mer RNA oligo is used as a control.

3.9. 3’-Adapter ligation

  1. Centrifuge “footprint” and “mRNA” samples at 20,000 x g for 30 min at 4°C, remove the supernatant. Centrifuge at 20,000 x g for 30 sec, remove the rest of the supernatant with a gel-loading tip, and air-dry the pellet for 10 min.

  2. Resuspend the pellet in 4.75 μL of RNase-free water. Add 2 μL of 50% PEG-8000, 1 μL of 10X T4 RNA ligase buffer, 1 μL of 3’-adapter, 0.25 μL of Superase-In, 1 μL of T4 RNA ligase, pipette up and down to mix. Incubate overnight at 16°C.

  3. Remove the excess of the adapter by adding 0.5 μL of 5’-deadenylase and 0.5 μL of Rec J exonuclease to the ligation reaction. Incubate at 30°C for 30 min.

  4. Add 30 μL of RNase-free water, 4 μL of 3M NaOAc, 1 μL of glycogen, and 2.5 volumes of 100% ethanol. Incubate at −20°C for at least 1 hr to precipitate RNA.

3.10. Reverse Transcription

  1. Centrifuge “footprint” and “mRNA” samples at 20,000 x g for 30 min at 4°C, remove the supernatant. Centrifuge at 20,000 x g for 30 sec, remove the rest of the supernatant with a gel-loading tip, and air-dry the pellet for 10 min.

  2. Resuspend the pellet in 11.5 μL of RNase-free water. Add 0.5 μL of 8 μM RT primer and 1 μL of 10 mM dNTPs. Incubate at 65°C for 5 min, put on ice.

  3. Add 4 μL of 5X FS buffer (supplied with Superscript III reverse transcriptase), 2 μL of 0.1 M DTT, 0.5 μL of Superase-In, 0.5 μL of Superscript III reverse transcriptase. Incubate for 30 min at 48°C, 1 min at 65°C, 5 min at 80°C.

  4. Hydrolyze RNA, by adding 0.8 μL of 2M NaOH, incubate at 98°C for 30 min. Add 0.8 μL 2M HCl to neutralize reaction.

  5. Add 20 μL of RNase-free water, 4 μL of 3M NaOAc, 1 μL of glycogen, and 2.5 volumes of 100% ethanol. Incubate at −20°C for at least 1 hr to precipitate DNA.

  6. Centrifuge the samples at 20,000 x g for 30 min at 4°C to pellet the DNA, remove the supernatant. Centrifuge at 20,000 x g for 30 sec, remove the rest of the supernatant with a gel-loading tip, and air-dry the pellet for 10 min.

  7. Resuspend the pellet in 5 μL of RNase-free water. Add 5 μL of 2X TBE-urea sample buffer.

  8. Prepare oligo sizing control containing 1 μL of 2.5 μM RT primer, 1 μL of 2.5 μM 128 nt marker oligonucleotide, 3 μL water, and 5 μL of 2X TBE-urea sample buffer.

  9. Heat the samples at 75°C for 3 min, spin down at max speed for 10 sec, put on ice for 1 min.

  10. Rinse wells of the 10% TBE-urea gel from urea and pre-run the gel at 180 V for 15 min in 1X TBE buffer.

  11. Load each sample into 1 well of the 10% TBE-urea gel, run the gel at 180 V for 50 min.

  12. Dissolve 5 μL of SYBR Gold in 50 mL of RNase-free water. Stain the gel with SYBR Gold for 5 min, protect from light.

  13. Using a blue light transilluminator, cut the gel slices around 128 nt and higher with a razor blade (Fig. 4). Freeze the polyacrylamide gel slices at −80°C for 10 min.

  14. Heat the gel slices at 70°C for 2 min, grind the gel using RNase-free disposable pellet pestles. Add 300 μL of 20 mM Tris-HCl pH 7.0, and incubate at 37°C for 3 hrs to extract DNA.

  15. Centrifuge the samples at 12,000 x g for 5 min at 4°C and transfer the supernatant to a new 1.5 mL tube. Add 30 μL of 3 M NaOAc,3 μL of glycogen, and 2.5 volumes of 100% ethanol. Incubate at −20°C for at least 1 hr to precipitate DNA.

Fig. 4.

Fig. 4

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Representative size selection gels used to isolate footprint and mRNA samples after reverse transcription. (a) Footprint samples obtained after reverse transcription were separated on 10% polyacrylamide TBU-urea gel. A mixture of RT primer and 128 nt marker oligonucleotide is used as a control. Cut the gel slice around 128 nt (upper band), which corresponds to the reverse transcription product. (b) For mRNA samples, cut just above the 128 nt marker. The size of the excised product should be around 150–170 nt.

3.11. Circularization

  1. Centrifuge the samples at 20,000 x g for 30 min at 4°C, remove the supernatant. Centrifuge at 20,000 x g for 30 sec, remove the rest of the supernatant with a gel-loading tip, and air-dry the pellet for 10 min.

  2. Resuspend the pellet in 16.75 μL of RNase-free water. Add 2 μL of 10X ssDNA ligase buffer (supplied with enzyme), 1 μL of 50 mM MnCl2, 0.25 μL of ssDNA ligase. Incubate at 60°C for 1 hr, immediately heat at 80°C for 10 min to inactivate the enzyme. Store the ssDNA ligation reaction product at −20°C.

3.12. PCR Library Amplification

  1. Set up PCR reactions on ice by mixing the following in a PCR tube: 146 μL of RNase-free water, 2 μL of 20 μM Forward primer, 2 μL of 20 μM Indexed Reverse PCR primer, 40 μL of 5X Phusion high-fidelity DNA polymerase buffer, 4 μL of 10 mM dNTPs, 4 μL of ssDNA ligation reaction product, and 2 μL of Phusion high-fidelity DNA polymerase. Choose Indexed Reverse PCR primer specific for each sample. Pipette 50 μL of the PCR mixture into 4 different PCR tubes.

  2. Perform the PCR amplification with varying number of cycles (6, 8, 10, 12) using the following settings:
    • Initial denaturation: 1 min at 98°C
    • Denaturation:15 sec at 94°C
    • Annealing: 5 sec at 55°C
    • Elongation: 10 sec at 65°C
    • Final extension: 2 min at 65°C

  3. To each PCR product add 5 μL of 3M NaOAc, 1 μL of glycogen, and 2.5 volumes of 100% ethanol. Incubate at −20°C for at least 1 hr to precipitate DNA.

  4. Centrifuge the samples at 20,000 x g for 30 min at 4°C to pellet the DNA, remove the supernatant. Centrifuge at 20,000 x g for 30 sec, remove the rest of the supernatant with a gel-loading tip, and air-dry the pellet for 10 min.

  5. Resuspend the pellet in 8 μL of RNase-free water, add 2 μL of non-denaturing 5X DNA loading dye.

  6. As a control, prepare a sample containing 0.5 μL of 10 bp DNA ladder, 7.5 μL water, and 2 μL of non-denaturing 5X DNA loading dye.

  7. Pre-run the non-denaturing 8% TBE gel at 180 V for 15 min in 1X TBE buffer.

  8. Load each sample into 1 well of the non-denaturing 8% TBE gel, run the gel at 180 V for 35 min.

  9. Dissolve 5 μL of SYBR Gold in 50 mL of RNase-free water. Stain the gel with SYBR Gold for 5 min, protect from light.

  10. Using a blue light transilluminator, cut the gel slices around 150 bp for “footprint” samples and around 180 bp for “mRNA” samples (Fig. 5) with a razor blade. Freeze the polyacrylamide gel slices at −80°C for 10 min.

  11. Heat the gel slices at 70°C for 2 min, grind the gel using RNase-free disposable pellet pestles. Add 300 μL of 20 mM Tris-HCl pH 7.0, and incubate at 37°C for 3 hrs to extract DNA.

  12. Centrifuge the samples at 12,000 x g for 5 min at 4°C and transfer the supernatant to a new 1.5 mL tube. Add 30 μL of 3 M NaOAc,3 μL of glycogen, and 2.5 volumes of 100% ethanol. Incubate at −20°C for at least 1 hr to precipitate DNA.

  13. Centrifuge the samples at 20,000 x g for 30 min at 4°C, remove the supernatant. Centrifuge at 20,000 x g for 30 sec, remove the rest of the supernatant with a gel-loading tip, and air-dry the pellet for 10 min.

  14. Resuspend the amplified sequencing library in 20 μL of RNase-free water. Proceed to Library quantification and High-throughput Sequencing.

Fig. 5.

Fig. 5

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Representative size selection gels used to isolate PCR amplified sequencing libraries. Following PCR amplification with varying number of cycles, the samples were separated on non-denaturing 8% polyacrylamide TBE gels. Select the desired number of cycles, in which the PCR products form a single, but bright enough band. Avoid lanes with high background. Cut the band ~150 bp for footprint libraries, and 170–190 bp for mRNA libraries.

3.13. Library Quantification and High-throughput Sequencing

  1. The quantity and the size distribution of the amplified sequencing library can be determined using the Bioanalyzer High Sensitivity DNA assay. The expected size is 148–152 bp for “footprint” library, and 170–190 bp for “mRNA” library [9].

  2. We usually multiplex 12 samples labeled with individual barcodes in a single sequencing run. For multiplexing, perform accurate quantification of the libraries using a qPCR-based sequencing library quantification assay. Mix the libraries in equimolar ratios to achieve 10 nM final concentration of the pool. Store at −20°C.

  3. Sequence the pooled libraries using a single-end 50 bp run on an Illumina platform. Detailed instructions for downstream analysis of the Ribo-Seq data have been described previously [17]. For normalization of the Ribo-Seq sequencing data in young and old cells using an internal spike-in see Note 12.

4. Notes

  1. Handle biotin-labeled cells gently, do not vortex.

  2. From this step adding antibiotics is not necessary. Extra care and aseptic techniques should be used to avoid contamination.

  3. It is important that the OD600 of the cell culture does not exceed 1.0. We noticed that the efficiency of the mother enrichment program is decreased with time. Fresh estradiol needs to be added after 20 h incubation.

  4. Sorting of cells should be performed on ice or in a cold room. Warming up the samples could cause the Dynabeads to lose biotin-binding capacity.

  5. We typically recover ~ 50% of the starting number of cells. The population of old cells obtained after 30 h incubation on average contains ~11–13 more bud scars per cell compared to the young population.

  6. It is important to always keep the sample frozen. Chill the tube in liquid nitrogen at least 10 sec between each grinding cycle. If necessary, combine several replicates to achieve 1×108 cells per sample that will be divided equally between ribosomal footprint and mRNA libraries. We also found that adding flash-frozen droplets of Lysis Buffer to the sample makes grinding more efficient.

  7. The amount of spike-in required for each sample is calculated based on OD260 values of the yeast lysates so that an equal amount of the worm lysate spike-in control (1%) is added into each sample proportional to the total RNA concentration. Alternatively, the same amount of the worm lysate spike-in could be added proportional to the cell number measurements (Section 3.2 step 25) to allow normalization on a per-cell basis.

  8. Other gradient makers or alternative methods for preparation of sucrose gradients can be used.

  9. If fewer than six samples are being analyzed, attach all buckets, and arrange the filled tubes symmetrically in the rotor. Opposing tubes must be filled to the same level with the 10–50% sucrose gradient.

  10. Once finished, clean the gradient fractionation system with RNase-free water. Thoroughly wash tubing and all removable components with warm water.

  11. Caution: acid-phenol:chloroform is toxic, avoid contact with skin and inhalation.

  12. For normalization of the Ribo-Seq data in young and old cells based on overall changes in protein translation, we utilize internal controls using spike-in with worm lysates. Following demultiplexing and trimming of the 3’ adapter sequence AGATCGGAAGAGCACACGTCT using Cutadapt software [18], sequencing reads are aligned to S. cerevisiae rRNA, tRNA, and sequences corresponding to C. elegans transcripts using Bowtie [19]. Reads that do not align are then mapped to the S. cerevisiae genome, and the number of read counts per gene is determined by HTseq-count software [20]. To normalize the translation between the libraries, yeast RPKM (reads per kilobase per million mapped reads) values are adjusted using a global linear scaling factor calculated by linear regression of worm spike-in RPKM values between the libraries [21,22].

Acknowledgments

This work was supported by the National Institutes of Health grants AG054566 and AG058713 to VML.

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