Rapid In Vivo Fixation and Isolation of Translational Complexes from Eukaryotic Cells

1. Yeast cell protocol

1. Yeast cell culture and fixation
   NOTE: Cell fixation and harvesting are adapted from^10,38 with modifications.
   1. Set up 1 L yeast cell culture (wild-type (WT) BY4741 are given as an example) in an orbital shaker with the starting optical density of no more than 0.05 AU at 600 nm (OD₆₀₀) in suitable media (1% w/v of yeast extract, 2% w/v of peptone, 2% w/v of dextrose (glucose), 40 mg/L of adenine sulfate (YPD) used as an example) under the desired conditions (30 °C used in this experiment).
   2. Set up a preparative centrifuge with compatible rotor and centrifuge bottles for pelleting the liquid suspension culture of yeast cells. For glucose starvation experiments, pellet the cells once the optical density of 0.6-0.8 AU at 600 nm (OD₆₀₀) is reached, using a brief centrifugation at 30 °C, 5,000 x g for 1 min.
      NOTE: Keep record of the OD of the growing cells and let the cells grow until the OD₆₀₀ reaches 0.6-0.8 AU, if the exponential growth phase is of an interest.
   3. Resuspend the pellet immediately in warm (30 °C) YP media containing no or low (0.25% w/v) added glucose and incubate the culture for a further 10 min at 30 °C in an orbital shaker-incubator.
      NOTE: Media composition might affect subsequent crosslinking efficiency. This protocol was tested using YPD only. When performing starvation experiments, adhering to the timing and minimizing the delays between procedures is critical.
   4. Once the cells are ready, set up an ice box inside the fume hood with a beaker containing 250 g of clean crushed water ice. Ensure 25 mL stripettes and freshly purchased methanol-stabilized 37% w/v formaldehyde solution are accessible inside the hood. Pour the 1 L culture into the beaker containing 25% w/v of crushed water ice.
      NOTE: Keep the cells on ice throughout all subsequent operations until the cells are frozen, unless indicated otherwise.
   5. Add 75 mL of 37% w/v of formaldehyde solution to a final concentration of 2.2% w/v and intensely stir the mixture until the ice melts.
   6. Once the ice is melted, set up a timer for 10 min.
      NOTE: Adhere to the recommended timings and temperature regimen to attain reproducible fixation results.
   7. After incubating for 10 min, transfer the culture into the precooled centrifuge bottles and pellet the cells by centrifugation at 4 °C, 5,000 x g for 5 min. While this spin is on, precool a 50 mL tube and keep freshly prepared buffer A (containing glycine to neutralize any remaining formaldehyde) on ice.
      NOTE: Refer to the supplied table for the exact buffer compositions.
   8. After centrifuging, place the centrifuge tubes onto ice with the pellet side in contact with the ice. Bring the tubes into the fume hood and discard the supernatant into a formaldehyde waste container.
   9. Resuspend the cell pellet from all tubes in 20 mL of buffer A using a 25 mL stripette and transferring to a 50 mL tube.
      NOTE: This wash is critical for avoiding irreproducible crosslinking and the buffer addition must not exceed 20 min of time from the cells' harvest.
   10. Make the volume up to 40 mL with buffer A and collect the washed cells by centrifugation at 4 °C, 5,000 x g for 5 min.
   11. Discard the supernatant and resuspend the cell pellet in 40 mL of buffer A1, which is buffer A not containing glycine, to remove any glycine contamination.
   12. Pellet cells again by centrifugation at 4 °C, 5,000 x g for 5 min.
   13. Repeat the washes with buffer A1 one more time. Discard the supernatant and place the cell pellet on ice. Weigh the tube with the pellet (wet cell mass should be ~1 g per 1 L of the cell culture).
2. Yeast cell disruption and cytosol collection
   1. Fill a polystyrene foam box lined with aluminum foil with liquid nitrogen to a depth of approximately 3 cm. Place a 50 mL tube upright in the box.
   2. Resuspend the pellet (~1 g wet cell mass) in 550 µL of buffer A2 by pipetting and vortexing for 10 s. Add 10 µL of 40 U/µL of RNase inhibitor and vortex again for 10 s.
      CAUTION: Wear appropriate protective equipment, such as thermally insulated gloves, when handling liquid nitrogen. Ensure that any container used to hold liquid nitrogen does not leak, and that the tube rack inside will not float up or fall on its side. Work in a well-ventilated area to avoid oxygen depletion.
   3. Using a 1 mL pipette, drip the cell suspension into the 50 mL tube containing the liquid nitrogen.
      NOTE: The dripping must be performed slowly and carefully to avoid aggregation of the droplets. Ensure the droplets freeze before introducing new droplets.
   4. Transfer the 50 mL tube with the frozen cell suspension droplets to room temperature and wait until the liquid nitrogen evaporates completely. Seal the tube with its cap and store the cell pellets at -80 °C or immediately proceed further.
      CAUTION: Ensure that the liquid nitrogen is completely evaporated before sealing the tube. Leftover liquid nitrogen in a sealed tube can cause a hazardous pressure build-up.
   5. To prepare for the next step, precool 1.5 mL nuclease-free tubes and 10 mL stainless steel grinding jars on dry ice.
   6. Transfer the frozen cell suspension droplets into the jars using a clean, sterile spatula.
      CAUTION: Ensure that the grinding jars are tightly sealed.
   7. Submerge the grinding jars into the liquid nitrogen for 1 min ensuring the liquid phase remains below the junction. Set up a cryo mixer mill at 27 Hz for agitation for 1 min.
      NOTE: Always balance the grinding canister with another one of the same model even if the sample requires only one canister for processing.
   8. Agitate the sealed grinding jars at 27 Hz for 1 min in the mixer mill.
   9. Re-cool the grinding jars in liquid nitrogen as before and shake at 27 Hz for 1 min further in the mixer mill.
   10. Transfer the jars to the ice box containing dry ice along with the 1.5 mL nuclease-free tubes. Using a small steel spatula, transfer the resultant powdered sample into the tubes in ~100 mg aliquots, and store the tubes at -80 °C.
       ​NOTE: It is recommended to use ~600 mg of the sample per experiment comprising polysome sedimentation profile analysis, separation of the cytosol into translated and non-translated fractions, and further separation of the translated fraction into SSU, ribosome, and disome fractions upon RNase digestion.
3. Separation of the fixed (poly)ribosomal complexes from the non-translated fractions of the cytosol
   NOTE: The procedure established earlier^10,38 is generally followed to enrich translated RNA based on its co-sedimentation with (poly)ribosomes. A more refined approach to separating the translated and non-translated cytosol fractions is introduced here, eliminating the need to precipitate and subsequently re-solubilize the material.
   1. Prepare 2.5 mL linear 10%-20% w/v sucrose gradients with buffer B using the freeze-thaw method⁴³ in thin wall ultracentrifuge tubes (5 mL, 13 x 51 mm).
      NOTE: The freeze-thaw method is performed by the sequential addition and freezing of buffered sucrose layers with linearly regressing concentrations on top of each other. See Supplementary Table 1 for details.
   2. To create a discontinuous 50% w/v sucrose cushion, upon the linear gradients thawing and stabilizing, slowly dispense 0.5 mL of 50% sucrose in buffer B directly onto the bottom of the tubes using a 1 mL syringe attached to 19 G x 1.5" needle or a glass capillary of similar/suitable dimensions. Before dispensing, carefully and slowly drive the tip of the needle or capillary from the top to bottom of the pre-formed sucrose gradients, avoiding any disturbance, until it reaches the tube bottom.
      NOTE: See Supplementary Table 1 for instructions on preparation of buffer B.
   3. Carefully balance the gradients by removing the top portions or layering more 10 w/v of sucrose in buffer B and keep them ice-cold or at 4 °C.
      NOTE: The discontinuous gradient with the bottom 50% sucrose layer is needed to collect material with higher sedimentation rate without precipitating it on the tube wall.
   4. Thaw ~100 mg of the frozen cell powdered sample at room temperature and immediately place on ice. Mix in 150 µL of buffer A2 by pipetting, add RNase inhibitor to 1 U/µL and mix by vortexing (avoid excessive foaming and mixing with the gaseous phase) for 10 s.
      NOTE: Continue all operations while keeping the material on ice, unless otherwise indicated.
   5. Pellet the cell debris by centrifuging the tubes at 4 °C, 13,000 x g for 5 min and recover the clarified supernatant (~150 µL) in a new 1.5 mL low protein binding tube.
   6. Load the resultant clarified mixture onto the discontinuous sucrose gradient tubes from step 1.3.3 and carefully balance them.
   7. Ultracentrifuge the tubes in a medium volume swing-bucket rotor at 4 °C, with average g-force 287,980 x g (k-factor 49) for 1 h 30 min.
      NOTE: These conditions have been pre-optimized (using post-ultracentrifugation gradient UV absorbance trace analysis) to retain the free (non-(poly)ribosomal) SSUs and LSUs (large ribosomal subunit) in the top (10%-20% sucrose) portion of the gradient while concentrating the (poly)ribosomal fraction in the bottom (50%) sucrose cushion without pelleting the material.
   8. Use a new sterile 1 mL syringe equipped with a 19 G x 1.5" needle to collect the translated cytosol fraction. Place the 5 mL gradient on a stable rack ensuring the bottom of the tube is visible.
   9. From the top of the tube, stick the needle straight into the bottom of the gradient (without puncturing the tube) and gently, without creating any bubbles, draw exactly 0.5 mL of the bottom solution containing the translated RNA pool.
      NOTE: Ensure this step is performed in a cold room and the tube is held firmly. It is recommended to draw the entire 0.5 mL in a single upstroke motion to avoid disturbance of the gradient.
   10. Confirm the (poly)ribosomal presence and the depletion of the SSU, LSU and lighter fractions in the resultant mixture by absorbance readout of the sucrose gradient upon ultracentrifugation run.
   11. Concentrate the collected translated RNA pool from the previous step to 100 µL using ultrafiltration in a micro-concentration device with 10 kDa cut-off regenerated cellulose membrane.
       NOTE: Pre-wash the micro-concentration device's membrane with 0.5 mL of buffer 1 (see Figure 2a) and use spin conditions (g) recommended by the manufacturer.
   12. Further dilute the material from the previous step five times (add 400 µL) with buffer 1 and concentrate back to 200 µL, to allow for a smaller volume as well as partial removal of the sucrose.
       NOTE: It is recommended to store the resultant mixtures at -80 °C for up to 6 months and use as an input material for the 'total translated RNA' RNA-seq library construction, or the RNase digestion step of the TCP-seq library construction. The 'non-translated' cytosol fraction can be recovered from the top of the gradient using a similar procedure and stored at -80 °C.
4. RNase digestion of the fixed (poly)ribosomal complexes and separation of digested material into small ribosomal subunit (SSU), monoribosomal (ribosomes, RS), and diribosomal (disomes, DS) fractions
   NOTE: The procedure generally follows an approach described previously^10,38 but a modified gradient type, separation time, acceleration and RNase digestion conditions are employed, to achieve best resolution across all three isolated fractions.
   1. Prepare carefully balanced 12.5 mL linear 10%-40% w/v sucrose gradients made with buffer 1 in 13 mL thin wall polypropylene tubes, 14 x 89 mm, using the freeze-thaw method⁴³ as described in step 1.3.1 and note therein.
   2. Thaw at room temperature and immediately transfer the samples on ice or take the concentrated and sucrose-depleted translated cytosol fraction from step 1.3.12.
      NOTE: Continue all procedures on ice unless otherwise indicated.
   3. Digest the translated cytosol fraction by mixing in 4.5 U of E. coli RNase I per 1 OD₂₆₀ unit of the fraction for 30 min at 23 °C. Immediately add and mix in by pipetting the RNase inhibitor capable of inactivating RNase I to 0.25 U/µL to the mixture, to inactivate RNase I.
      NOTE: Use RNase inhibitor capable of inhibiting RNase I. Derive AU₂₆₀ by using AU₂₆₀ = (Absorbance at 260 nm standardized to optical density units equivalent to 1 cm optical path x volume of the lysate in µL) / 1,000.
   4. Immediately transfer the samples to ice.
      CAUTION: It is critical to adhere to the recommended conditions of digestion and carefully measure the amount of the added RNase I. The RNase I unit referred to here is defined as the amount of the enzyme required to produce 1 µg of acid-soluble material from mouse liver RNA in 30 min at 37 °C. RNase I batches may have undocumented variations in activity and may require experimentation to achieve optimal digestion conditions. If the enzyme stock is too concentrated, it is recommended to dilute it with buffer 1 to avoid pipetting very small volumes of the solution.
   5. Load the reaction mixtures onto the 10%-40% w/v sucrose gradients from step 1.4.1.
      NOTE: Use final volumes in the range of 150-300 µL per gradient. Each purification requires minimally two gradients. Use different input volumes of the material (lower AU₂₆₀, 10-11 AU₂₆₀, for DS and comparatively higher AU₂₆₀, 13-14 AU₂₆₀, for SSU or RS) to achieve optimal separation.
   6. Ultracentrifuge the tubes in a medium volume swing-bucket rotor at 4 °C with average g-force 178,305 x g (k-factor 143.9) for 3 h 30 min.
      CAUTION: If spare balance tubes are needed, equalize their mass and mass distribution with the sample-containing tubes. Use spare sucrose gradients overlaid with an amount of buffer equivalent to that of the sample overlay and not tubes with uniform sucrose concentration.
   7. Set up a gradient fractionator device at least 30 min before the ultracentrifugation spin completion, including filling in the 0.2 µm filtered heavy chase solution (e.g., 60% sucrose in deionized water as used here) into the displacement pump.
      NOTE: It is recommended to de-contaminate the lines and tubings of the fractionator using deionized water, followed by 1%-2% SDS solution in deionized water, deionized water, and finally 80% ethanol in deionized water solution before and after the runs.
   8. Adjust the absorbance readout baseline by first filling the system with deionized water and zeroing the optics as per the manufacturer's recommendations, and then compensating the baseline shift using a spare unloaded 14 x 89 mm sucrose gradient made with a buffer identical to the sample tubes (e.g., buffer 1).
      NOTE: Use the same displacement speed to make the adjustments as for the sample readout, such as 1.5 mL/min.
   9. Measure the displacement system dead volume by accurately counting time between the solution first entering the optical path of the detector and first appearing at the fraction collector output.
      NOTE: With the recommended speed of 1.5 mL/min, the fractionation can be performed at room temperature. It is recommended to immediately transfer the collected fractions on ice.
   10. Perform fractionation using live absorbance readout at 254 nm, 1.5 mL/min displacement speed, and in-line fraction detection based on the expected sedimentation position and absorbance profile of the samples. Use collector tube switching with a time delay corresponding to the dead volume as measured before.
   11. Isolate fractions corresponding to the positions and mobility of the SSU, RS, and DS complexes and collect them into new low protein binding 1.5 mL microcentrifuge tubes; immediately transfer the isolated fractions on ice and freeze if not processed further right away.
       NOTE: It is recommended to immediately flash-freeze the collected fractions in dry ice or liquid nitrogen and store at -80 °C or below for up to 6 months.
5. De-crosslinking of the ribosomal complexes and isolation of the RNA to construct RNA-seq libraries
   1. To de-block/reverse the crosslinks and isolate the RNA away from the associated proteins, transfer approximately half of the entire sucrose gradient fractions into new low nucleic acid binding nuclease-free polypropylene 1.5 mL microcentrifuge tubes (350 µL per tube) with lid safety/locking devices.
   2. Supplement the mixtures with 40 µL of 100% stop solution (10% SDS w/v and 100 mM EDTA), 4 µL of 1 M Tris-HCl pH 2 at 25 °C (to 10 mM), 1.6 µL of 2.5 M glycine (to 10 mM) and deionized nuclease-free water to obtain the final volume of 400 µL.
   3. Mix the contents of the tubes by pipetting and transfer the tubes at room temperature.
   4. Add equal volume of the acidic phenol:chloroform:isoamyl alcohol 125:24:1 (pH 4.0-5.0) mixture to each tube. Vigorously shake the mixtures for 2 min using a vortex mixer set to maximum speed.
      CAUTION: Phenol and chloroform are corrosive and toxic. Avoid physical contact with the liquids and work in a well-ventilated area or under a fume hood. Always use gloves, lab coat and protective goggles or a face shield when working with phenol or chloroform.
   5. Place the tubes in a thermoshaker and continuously shake at 65 °C, 1,400 rpm for 30 min.
   6. Facilitate phase aggregation by centrifuging the mixture at 12,000 x g for 10 min at room temperature.
   7. Collect the upper aqueous phases and transfer them into fresh low nucleic acid binding 1.5 mL tubes.
      NOTE: To avoid cross-contamination, do not attempt to recover the aqueous phases completely. A reasonable recovery volume is 300-350 µL.
   8. Supplement the collected aqueous phases with 0.1 volumes of 3 M sodium acetate (pH 5 at 25 °C), 20 µg of glycogen (using 5 µg/µL stock) and 2.5 volumes of absolute ethanol. Carefully mix the solutions by vortexing the tubes for 1 min.
   9. Precipitate the RNA by incubating the samples at -20 °C for at least 2 h (recommended overnight).
   10. Warm the tubes to room temperature and mix by vortexing.
       NOTE: Pre-warming of the tubes and subsequent centrifugation at room temperature (without forced chilling) help to reduce salt and phenol co-precipitation and carryover. These conditions should not result in material loss or inefficiency of RNA collection if performed as described and using sufficiently pure ethanol.
   11. Pellet the RNA precipitate by centrifuging the tubes at 12,000 x g for 30 min at room temperature.
   12. Discard the supernatant and wash the pellet twice with 80% v/v ethanol, collecting it each time by centrifugation at 12,000 x g for 10 min at room temperature.
   13. Dry the RNA pellets by opening the tube lids and placing the opened tubes in a dry-block heater set to 45 °C for 10 min. Dissolve the resultant dried pellet in 20 µL of 1x HE buffer.
   14. Estimate the resultant RNA concentration using UV absorbance spectrum measurement.
       NOTE: RNA fragment length and total amount can be further assessed using denaturing gel-electrophoresis, such as in an automated fluorescence-based capillary gel electrophoresis apparatus.
6. Selective co-immunopurification of the SSUs by the tagged eIFs and western blot analysis of the selective SSU enrichment
   ​NOTE: Use ~15 AU (260 nm) of the digested and sedimentation-segregated SSU fraction from step 1.4.11 to perform affinity purification using magnetic IgG beads. Save ~5% of the SSU fraction as input control (Input fraction, I). eIF4A-tagged (TIF1-TAP; Tandem Affinity Purification tag) yeast strain was used which also makes it possible to detect eIF4A by probing for the TAP-tag using anti-TAP antibody.
   1. Transfer 100 µL of magnetic IgG beads suspension (1 mg of the beads were used for each 15 AU (260 nm) of the lysate or fraction) into a new low protein binding 1.5 mL tube; collect the beads using magnetic rack and aspirate them.
   2. Wash the magnetic beads twice with 1 mL of buffer 1 by using sequential resuspension by pipetting and collection using the magnetic rack.
   3. After washing, collect and decant the beads, while keeping them on the magnetic rack.
   4. Add the SSU fraction to the washed beads and incubate the mixture for 4 h with rotation at 4 °C in a cyclomixer set at ~20 rpm.
   5. Collect the beads using the magnetic rack at 4 °C and save the supernatant (Flow-through fraction, FT).
   6. Wash the beads twice at 4 °C with buffer 1 supplemented with 4 mM DTT, each time rotating for 10 min in the cyclomixer and collecting and decanting the beads on the magnetic rack. Save the washes (W1 and W2 fractions).
   7. For an analytical application such as western blotting, elute the bound material under denaturing and reducing conditions by adding LDS (lithium dodecyl sulfate) polyacrylamide gel electrophoresis (PAGE) sample buffer with pH 8.5 to 1x and DTT to 2 mM.
   8. Heat the mixture at 95 °C for 5 min in a thermal block to finalize the elution.
   9. Collect the beads using the magnetic rack and recover the denatured eluate (E fraction) in a fresh low protein binding 1.5 mL microcentrifuge tube.
   10. Use the E fraction from the previous step to run a denaturing sodium dodecyl sulfate (SDS) PAGE immediately, or store the E fraction at -20 °C.
       NOTE: For a preparative collection of the TAP-tag-enriched translational complexes for any subsequent application, use an alternative elution approach employing Tobacco Etch Virus (TEV) protease. Refer to the Supplementary Table 1 for further details.
   11. To concentrate the dilute FT, W1 and W2 fractions, precipitate their material by adding 3x volumes of ice-cold acetone. Incubate the sample-acetone mix at -20 °C for 3 h.
   12. Pellet the precipitate by centrifuging the tubes at 13,000 x g for 10 min at 4 °C.
   13. Discard the supernatant and air dry the pellet in the open tubes at room temperature for 30 min.
   14. Dissolve the pellet in 7 µL of 1x LDS loading buffer supplemented with 2 mM DTT. Heat the samples in a thermal block set to 95 °C for 5 min.
   15. Load all I, FT, W1, W2, and E samples onto a 4%-12% w/v of acrylamide gradient, Bis-Tris polyacrylamide denaturing gel. Run the gel using 1x MES SDS (2- [N-mopholino]ethanesulfonic acid, sodium dodecyl sulfate) running buffer at 80 V, until the protein marker (10-250 kDa) resolves well and the lead dye reaches the bottom of the gel.
       NOTE: It is recommended to load serial dilutions of the WCL (whole cell lysate) (2-10 µg) on the gel as a control. It may take several attempts to achieve comparable loading of the gel across the fraction material.
   16. Transfer the protein content of the gel onto a polyvinylidene difluoride (PVDF) membrane by wet transfer method at 100 V for 1 h in a cold room as recommended by the western-blotting equipment manufacturer.
   17. Block the membrane using an appropriate blocking buffer (Phosphate Buffered Saline based) at room temperature for 1 h under constant shaking.
   18. Following manufacturer's instructions for antibody dilution, probe the membrane with anti-TAP-antibody for detecting the tagged eIF4A protein, anti-Pab1p antibody or anti-β-actin antibody (or any other desirable target) by overnight incubation of the membrane with Blocking Buffer (PBS)-diluted antibody (1:1,000 dilution) in a cyclomixer in a cold room.
       NOTE: 1:1,000 antibody dilution is a good starting point.
   19. Wash the membrane three times with 1x Phosphate Buffered Saline, 0.2% v/v Tween 20 (PBST) for 10 min each.
   20. Probe the membrane with fluorescently labeled secondary antibodies following manufacturer's instructions by incubating in a cyclomixer at room temperature for 1 h.
       NOTE: 1:20,000 antibody dilution is a good starting point.
   21. Wash the membrane three times with 1x PBST for 10 min each. Briefly rinse the membrane with deionized water, and then with absolute methanol. Dry and visualize the membrane in a fluorescent imaging system according to the manufacturer's instructions.
       ​NOTE: Staining for other proteins can be achieved by using secondary antibodies with dyes matching different fluorescent channels (such as in the eIF4A-TAP vs. β-actin pair used here), by sequential staining or stripping and staining the same membrane or cutting the membrane from a gel loaded with repeating pattern of fractions and separately probing each piece with respective antibodies (as in the Pab1p example used here).

2. Mammalian cell protocol

1. Mammalian cell culture and fixation
   1. In 2 T-175 flasks, grow HEK293 cells to 60%-70% confluence in Dulbecco's Modified Eagle Medium and 10% v/v Fetal Bovine Serum at 37 °C and 5% v/v carbon dioxide.
      NOTE: The complete media is made by adding 55 mL of commercial FBS into a 500 mL of commercially purchased DMEM with high glucose, containing L-glutamine, phenol red and sodium bicarbonate, but no HEPES or sodium pyruvate. Cell counts per T-175 flask at 70% confluence should be in the range of 1.7-2.0 x 10⁷.
   2. At least 3 h prior to the desired fixation time, replace the media of the T-175 flasks with precisely 30 mL of pre-warmed complete media and replace the flasks in a cell incubator.
      NOTE: Ensure that the fresh media is pipetted onto the opposite side of the flask to the cell monolayer to avoid cell detachment. Attempt to conduct the media exchange as quickly as possible, introducing minimal gas and temperature balance disturbance.
   3. Once the cell media has been replaced, prepare buffers and chemicals required for fixation. Prepare Dulbecco's Phosphate Buffered-Saline (DPBS) with 50 mM glycine by adding 10.2 mL of 2.5 M glycine stock to a 500 mL bottle of DPBS and mixing.
   4. Prepare a bottle of DMEM supplemented with 10% FBS as in step 2.1.1 to be used in non-sterile conditions and a 100 mL aliquot of 0.25% Trypsin-EDTA. Source an additional bottle of commercial DPBS pre-formulated with calcium chloride (CaCl₂) and magnesium chloride (MgCl₂).
      NOTE: The solutions can be stored at 4 °C for up to 2 weeks.
   5. Prepare an ice box to the brim with crushed water ice such that a T-175 flask can fit evenly on top and keep in the fume hood along with the prepared buffers, also on ice.
      NOTE: Due to the rapid responses of translation to any environmental change, all timings between the removal of the cell flasks from the incubator and addition of the formaldehyde solution must be minimized.
   6. To snap chill the cells, remove the T-175 flask from the incubator and firmly press it against the ice ensuring maximal surface contact. Inside the chemical fume hood, tilt the flask onto its side so that the media collects at the side opposite to the cells. Pipette 168 µL of 37% w/v formaldehyde directly into the pooled media (to a final concentration of 0.2% w/v). Immediately mix by gently rocking the flask back and forth, close and reposition the flask on ice, ensuring it is horizontal and the cells are covered evenly.
      CAUTION: Formaldehyde is a harmful substance with potential long-term adverse effects and also an irritant to both the respiratory system and skin. It should only be handled in a suitable chemical fume hood. Containers of formaldehyde must always be sealed when outside of the fume hood.
      NOTE: Ensure that the formaldehyde is added directly into the cell media and not to the flask wall. Step 2.1.6 should take less than 1 min.
   7. Incubate the flasks on ice for a further 10 min. Pour off the media into an appropriate waste container through the flask side opposite to the cells.
   8. Using a stripette, pipette in 30 mL of Dulbecco's Phosphate Buffered Saline without calcium and magnesium ions and additionally containing 50 mM glycine, gently on the side opposite to the cells. Mix by rocking the flask; return the flask to horizontal position and incubate for 10 min more on ice.
   9. Pour off the solution through the flask side opposite to the cells and gently add 7 mL of the standard 0.25% w/v Trypsin-EDTA solution to detach and resuspend the cells. Incubate the flask at room temperature for 5-10 min.
      NOTE: Ensure Trypsin-EDTA solution covers all the cells evenly. Use periodic gentle tilting and rocking to promote cell detachment.
   10. Relocate the flask vertically and using a stripette collect the detached cells by gently washing any remaining cells from the flask walls. Transfer the suspension into a 50 mL tube set on ice.
       NOTE: Fixed cells can become more fragile; do not pipette intensely or more than what is required to detach the cells from the flask wall.
   11. Immediately supplement the collected cell suspension with 20 mL of complete media (the non-sterile ice-cold media with 10% FBS) and mix by gently flipping the tube.
       NOTE: The complete cell culture media (including 10% FBS) is added to neutralize the trypsin, preventing further damage to the cell membranes and cell disintegration.
   12. Pellet the cells by centrifuging the tube at 100 x g for 5 min and 4 °C. Cell pellet must be clearly visible.
   13. Pour off the media and gently resuspend the cell pellet in 10 mL of ice-cold DPBS with Ca²⁺, Mg²⁺, and without glycine.
   14. Repeat step 2.1.12.
   15. Pour off the wash buffer and gently resuspend the cell pellet in 800 µL of ice-cold DPBS with Ca²⁺, Mg²⁺, without glycine, on ice. Transfer the resuspended cells into a new low protein binding 1.5 mL microcentrifuge tube.
   16. Centrifuge the tube at 100 x g for 3 min and 4 °C. Carefully discard the supernatant using a 1 mL pipettor. At this stage, the cell pellet can be frozen at -80 °C or proceed to the cell lysis step.
       NOTE: Frozen cell pellets can be stored at -80 °C up to 1 year. We found that cell pellet freezing facilitates subsequent lysis and recommend freezing even if longer term storage is not planned.
2. Mammalian cell disruption and cytosol collection
   1. In a biosafety cabinet, add 300 µL of the lysis buffer based on nonionic, nondenaturing detergent and 7 µL of 40 U/µL RNase inhibitor. Mix well by pipetting using a 1 mL tip.
   2. Carefully attach a 25 G needle to a 1-3 mL syringe and vigorously pipette the mixture, using at least seven slow upward intake and fast downward exhaust strokes.
   3. Discard the syringe and needle into a sharps bin and repeat the procedure using a 0.3 mL syringe equipped with a 31 G needle.
   4. Discard the syringe and needle into a sharps bin. Centrifuge the tubes at 4 °C, 12,000 x g for 5 min to pellet the cell debris.
   5. Transfer the supernatant into a new low protein binding 1.5 mL microcentrifuge tube. Store both, the cell debris (for control purposes) and the resultant clarified cell lysate at -80 °C.
      ​NOTE: Optical density of the lysate ranges between 25-30 AU₂₆₀ when two T-175 flasks are combined and the recommended volumes followed. The lysates and cell debris can be stored at -80 °C up to 1 year.
3. Separation of the fixed (poly)ribosomal complexes from the non-translated fractions of the cytosol
   1. Prepare linear 15%-45% w/v sucrose gradients in 13 mL thin wall polypropylene tubes, 14 x 89 mm, using freeze-thaw method generally as described in step 1.3.1 of the yeast protocol, but using buffer 2 (Figure 2a).
      NOTE: Thaw the gradients overnight in a cold room at 4 °C the night prior to the fractionation.
   2. Load 150-250 (maximally 300) µL of the cell lysate from the previous step 2.2.5 onto the balanced gradients. Store the remaining lysate at -80 °C and use for control purposes.
      ​NOTE: Here an example of sedimentation-based segregation into polysomal, ribosomal and 'free' SSU fractions is provided. Refer to the provided Supplementary Table 1 for an alternative approach.
   3. Ultracentrifuge the tubes in a medium volume swing-bucket rotor at 4 °C, average g-force 178,305 x g (k-factor 143.9) for 1 h 45 min.
   4. 30 min prior to the spin completion, set up and baseline the gradient fractionator, as described in the yeast protocol steps 1.4.7-1.4.9.
   5. Fractionate the gradients generally as described in the yeast protocol steps 1.4.10-1.4.11.
      ​NOTE: This step will separate polysomal, ribosomal and 'free' SSU fractions. Polysomal fractions may be used in polysome profiling experiments.
   6. Immediately transfer the collected fractions on ice and if not further processed, store at -80°C up to 6 months.
      NOTE: If the fraction collector tube change is synchronized with the on-line fraction identification and segregation, we recommend using up to 800 µL fractions (collection time of 32 s per fraction at 1.5 mL/min). If the fractionation is performed without using the in-line absorbance readout, it is recommended using 250-500 µL fractions (10-20 s per fraction at 1.5 mL/min). Following separation, the fractions can be used for immunopurification, electron microscopy, denaturing PAGE and western blotting straight away, or subjected to crosslink reversal for subsequent RNA and/or proteomics analyses.