Isolation of Subcellular Ribosome Subpopulations Based on Recombinant Peptide Tag-Specific Location-Restricted Illumination-Enhanced Biotinylation

摘要

AviTag-specific Location-restricted Illumination-enhanced Biotinylation (ALIBi) is a novel optogenetically activated method developed for labeling and affinity-purifying subcellular populations of ribosomes for downstream sequencing and proteomic analysis.

摘要

Growing evidence suggests that mRNA translation is a highly compartmentalized process within a cell, and that subcellular trafficking and localization of specific mRNAs is key to ensuring that proteins with compartment-specific functions are produced in the ideal milieu and in appropriate quantities. However, techniques for subcellular isolation and characterization of the ribosomes that translate these mRNAs have been limited to date, such that much remains unknown about how the composition of translational machinery contributes to regulation of localized mRNA translation.

Here, we demonstrate AviTag-specific Location-restricted Illumination-enhanced Biotinylation (ALIBi), a method that combines epitope tagging, a newly developed optogenetically activated split-biotin ligase, and affinity purification to rapidly and specifically label and isolate ribosomes localized to any subcellular compartment of interest. First, CRISPR/Cas9 editing is used to fuse an AviTag peptide, a tobacco etch virus (TEV) protease cleavage site, and a FLAG epitope tag to a ribosomal protein. The split biotin ligase, fused to an organelle-targeting domain, is expressed in this cell line and is inactive under normal biotin concentrations. Upon activation by supplemental biotin and blue light illumination, the ligase biotinylates AviTagged ribosomes in the immediate vicinity, allowing for affinity purification of biotinylated ribosomes and associated proteins and mRNAs on streptavidin-coated beads. Non-denaturing elution via TEV protease cleavage yields samples suitable for downstream characterization of core ribosomal proteins, ribosome-associated proteins, and ribosome-bound mRNAs via RNA sequencing or mass spectrometry proteomics.

In this protocol, we review design principles for fusing AviTag to a ribosomal protein and targeting the split biotin ligase enzyme to the organelle of interest. We demonstrate activation of the ALIBi system, cell lysis, affinity purification, and sample elution. Finally, we discuss typical results and troubleshooting.

引言

There is an important need for tools that can isolate biological samples from specific subcellular compartments to better investigate how subcellular organization and trafficking factor into cellular processes. mRNA translation is a key example of such a process: growing evidence suggests that translation is highly compartmentalized and, in particular, that many mRNAs are localized to the organelles where their encoded proteins will reside^1,2. However, less is known about the subcellular localization of translational machinery. Ribosomes begin their existence in the nucleolus and nucleus, where assembly from ribosomal proteins (RPs) and ribosomal RNAs (rRNAs) begins. They are then exported to the cytoplasm for the final stages of assembly after which they begin translating mRNAs^3,4. Canonical sites of translation include the cytoplasm and endoplasmic reticulum; the latter is where integral membrane proteins and proteins destined for membrane-bound organelles or extracellular secretion are synthesized⁵. More recently identified sites of localized translation include the mitochondria, peroxisome, and neuronal processes^{6,7,8,9,10,11,12}. Interestingly, evidence is also emerging that translational machinery features such as ribosome-associated proteins, post-translational modifications, rRNA modifications, and non-canonical stoichiometry of core ribosomal proteins are involved in regulatory mechanisms that govern translation of specific mRNAs and parameters such as translation fidelity and rate^{13,14,15,16,17,18,19,20}. For example, hundreds of proteins have been identified that interact with the ribosome and could potentially lead to differences in subcellular ribosome composition¹³. However, techniques for isolating subcellular ribosome populations have been limited to date; thus, much remains unknown about how the composition of translational machinery contributes to regulating localized mRNA translation. The advent of versatile methods for isolating subcellular ribosome populations, particularly those compatible with mammalian model systems, will allow for exploration of these translation machinery features at the subcellular level.

The earliest methods for isolating subcellular fractions containing ribosomes were based on principles of sequential lysis and centrifugation^21,22. Typically, these methods are limited to certain organelles and often require extensive processing times. A more recent method is based on the biotin ligase BirA, which specifically biotinylates an AviTag peptide substrate^10,23,24. The BirA can be genetically localized to the organelle of interest in cells where ribosomes have been tagged with AviTag. However, because BirA is active at physiologic concentrations of biotin, it is necessary to culture cells under conditions of biotin depletion in order to avoid constitutive and non-specific biotinylation of AviTag-bearing ribosomes^10,24. This limits the use of BirA in cell types that are sensitive to biotin depletion²⁵. Finally, promiscuous biotinylation methods have been developed that use engineered enzymes such as BioID, TurboID, and APEX^26,27,28. Upon induction, these enzymes produce a reactive biotin intermediate that covalently modifies any lysine residues within a radius of the enzyme. Promiscuous biotinylation methods are highly useful for mapping subcellular proteomes, but less informative for distinguishing proteins in the general vicinity of localized ribosomes as opposed to proteins that are in closer proximity with them.

Here, we demonstrate our recently developed method for labeling and isolating subcellular ribosome populations, called AviTag-specific Location-restricted Illumination-enhanced Biotinylation (ALIBi). This method leverages an engineered split BirA fused to optogenetic enhanced Magnet (eMag) switches (Figure 1A) that heterodimerize under blue light illumination²⁹; and a peptide tag consisting of FLAG epitope, tobacco etch virus (TEV) protease cleavage site, and AviTag (FTA) fused to the Rpl31 ribosomal protein (Figure 1B). The split enzyme can be genetically targeted to selected organelles, is inactive at physiologic concentrations of biotin, and can be reconstituted within minutes upon blue light illumination and the addition of supplemental biotin (Figure 1C-E). Subsequent cell lysis, affinity purification, and non-denaturing enzymatic elution via TEV protease cleavage yields ribosomes and co-purifying molecules such as mRNAs and ribosome-associated proteins, which are suitable for downstream analytic methods such as RNA sequencing and mass spectrometry proteomics (Figure 1A). Therefore, this method reveals not only ribosome composition but also the mRNAs to which ribosomes are bound. In a recent article describing the seminal application of ALIBi to characterize subcellular ribosome populations, we successfully targeted the split enzyme to the cytoplasm, nucleolus, nucleus, endoplasmic reticulum, plasma membrane, and mitochondria (Table 1)³⁰. The resulting comparison of translational machinery between these subcellular compartments led to novel and unexpected findings regarding translation at the endoplasmic reticulum and mitochondria, as well as potential leads for future investigation of ribosome biogenesis factors in the nucleus and nucleolus.

This protocol contains a detailed demonstration of transfection and activation of the ALIBi system, cell lysis, affinity purification, and sample elution. We also discuss how to verify successful affinity purification via western blot and make recommendations for performing downstream RNA sequencing or mass spectrometry (MS) proteomic analysis (see Supplemental File 1). This protocol is written for experiments using mouse embryonic stem cells (mESCs) expressing Rpl31 fused to FTA (L31-FTA) homozygously from the endogenous Rpl31 locus; this cell line is available from the authors upon request. The strategy for generating this cell line is described in Supplemental File 1 and in Supplemental Figure S1 and Supplemental Figure S2. Plasmids encoding the split enzymes targeted to several organelles are available on Addgene (catalog numbers #235608 to 235643). The design principles for targeting the split enzymes to these organelles is described in Supplemental File 1. The protocol assumes that each sample originates from mESCs grown in a 10 cm plate. We have found that the 10 cm scale yields sufficient material for mass spectrometry proteomics, while the 6-well scale is sufficient for western blots or RNA sequencing.

方案

1. mESC cell culture

NOTE: Prior to ALIBi activation and cell lysis, mESCs should be handled using standard sterile technique for cell culture. Centrifugation of intact mESCs should be performed at 200 × g for 3 min at room temperature unless otherwise specified. Prewarm media to 37 °C in a water bath before use unless otherwise specified. mESCs should be passaged into a fresh tissue culture plate that has been gelatinized overnight (see step 1.1).

1. The day prior to thawing L31-FTA mESCs, coat the bottom of a 10 cm plate with 5 mL of 0.1% gelatin. Incubate at 37 °C overnight.
2. Combine mESC media components (Table 2) and pass through a 0.2 µm filter. Store at 4 °C for up to 1 month.
3. On the day of thawing, warm a 5 mL aliquot of media for diluting the thawed cells and a 10 mL aliquot for plating them.
4. Retrieve a cryovial containing ~5 × 10⁶ L31-FTA mESCs from liquid nitrogen storage. Immediately thaw by swirling the vial bottom in a 37 °C water bath.
5. Gently mix 0.5–1 mL of media from the 5 mL aliquot with the cryovial contents as the vial capacity allows, then transfer the cryovial contents back to the 5 mL media aliquot.
6. Pellet the cells by centrifugation. Aspirate the supernatant.
7. Resuspend the cell pellet with the 10 mL media aliquot.
8. Aspirate the gelatin from the 10 cm plate and immediately transfer cell suspension into the plate. Incubate at 37 °C.
9. Every 24 h (except on passaging days), aspirate old media from the plate and replace with 10 mL of fresh media.
10. When colonies are 70%–80% confluent (typically 48 h after plating), aspirate the old media and rinse twice with 10 mL of Dulbecco’s phosphate-buffered saline (DPBS).
11. After aspirating the DPBS, coat the plate with 2 mL of 0.05% trypsin (Table 2). Incubate at 37 °C for 5 min.
12. Neutralize the trypsin by adding 4 mL of cold mESC media to the plate. Immediately use a 5 or 10 mL serological pipet to pipet up and down 15x in the plate to dissociate colonies into a single-cell suspension.
13. Transfer the suspension into a conical tube. Optionally, remove an aliquot to count cells using a hemocytometer.
14. Pellet the cells by centrifugation. Aspirate the supernatant.
15. Resuspend the cell pellet in fresh media. Plate 5 × 10⁶cells (typically 1:6 passaging ratio) in a fresh gelatinized 10 cm plate and incubate at 37 °C. Repeat steps 1.9–1.15 as needed until ready to proceed with transfection.
16. To make frozen cell stocks, trypsinize a plate of mESCs (as in steps 1.10–1.14). Resuspend the cell pellet in freezing media (250–500 µL per 5 × 10⁶ cells), aliquot into cryovials, and freeze slowly in a cell freezing container at -80 °C overnight before transferring to liquid nitrogen storage.

2. Transfection

NOTE: Perform transfection on a day when mESCs are ready for passaging. Plasmids may begin expressing as early as 3-5 h after transfection. Once expressing, we recommend handling cells in the dark and minimizing exposure to temperatures below 37 °C to prevent premature activation, since previous work has shown that Magnet optogenetic domains dimerize not only under blue light illumination but also upon exposure to cold³¹. Keep plates in a dedicated incubator or shield a shelf in a shared incubator from light with aluminum foil. When handling cells outside the incubator, turn off or block all room lights and windows beforehand, and use a red-light headlamp to see. Do not leave the incubator door open for longer than necessary. Take only small batches of plates out of the incubator at a time, and rest plates on an insulating surface (i.e., a clean styrofoam box lid) while handling to minimize contact with room-temperature work surfaces. Warm media fully before use. Do not inspect plates under a light-emitting microscope.

1. The day before transfection, gelatinize a 10 cm plate.
2. On the day of transfection, aspirate gelatin from the plate and add 7.5 mL of media to prewarm at 37 °C. Prewarm reduced-serum medium at 37 °C for resuspending cells.
3. Dilute 18.75 µL of the transfection reagent in 250 µL of reduced-serum medium. Tap gently to mix and incubate for 5 min at room temperature.
4. If co-expressing two plasmids encoding the N- and C-fragments, add 3.75 µg of each plasmid to 250 µL of reduced-serum medium. If only expressing one plasmid (i.e., a GFP negative control or a full-length BirA positive control), add 7.5 µg of the single plasmid instead.
5. Add the plasmid/reduced-serum medium mix to the transfection reagent/reduced-serum medium mix and tap/invert to mix. Incubate at room temperature for 15 min.
6. In the meantime, trypsinize and pellet mESCs (steps 1.10–1.14). Resuspend the pellet in warmed reduced-serum medium to a concentration of 10 × 10⁶ cells/mL.
7. Add 7.5 × 10⁶ (750 µL) resuspended cells to the plasmid/transfection reagent/reduced-serum medium mixture. Invert the tube to mix. Incubate for 10 min at RT, inverting the tubes occasionally to keep cells suspended.
8. Invert the tubes again briefly and add the mESC/plasmid/transfection reagent/reduced-serum medium mixture to the plate containing warmed media. Gently rock the plate to evenly distribute the cells.
9. Incubate the plate at 37 °C for 3–5 h, allowing the cells to adhere within this time.
   NOTE: From this point on, we recommend observing the above precautions against exposure to light and temperatures below 37 °C.
10. Gently aspirate the old media. Add fresh warmed media, pipetting down the side of the plate to avoid dislodging cells. Resume incubation at 37 °C.
11. If activating/lysing at 24 h after transfection, proceed to section 3. If planning to activate/lyse at 48 h after transfection, change media ~24 h after plating the transfected cells.

3. ALIBi activation and cell lysis

NOTE: Continue to handle cells/samples in the dark until the beginning of elution (step 5.4). Once cells have been chilled on ice (step 3.15), keep samples cold and use only chilled reagents until the beginning of elution. Ensure that working surfaces, equipment, reagents, and plastics are RNAse-free by using freshly opened items and wiping or rinsing hard surfaces with an RNAse decontaminating solution and 70% ethanol. Wear gloves whenever handling samples or equipment. For samples intended for MS proteomics, also minimize keratin contamination by keeping the work space dust-free; cover bare skin, and tie back long hair.

1. Prechill the centrifuge to 4 °C.
2. With lights off, move the transfected plates to a temporary 37 °C incubator so they will not be exposed to temperature fluctuation in the main incubator after the transilluminator is placed inside.
3. Wipe down the blue light transilluminator with 70% ethanol and connect it to power. Place the transilluminator in the main incubator and allow incubator conditions to re-equilibrate while continuing with the setup.
4. Warm an aliquot of media to be dosed with biotin and cycloheximide (CHX) (10 mL of media per 10 cm plate) to 37 °C. If running control samples without supplemental biotin, also warm a separate aliquot to be dosed with CHX only.
   NOTE: Some buffer components can be mixed the day before use and stored overnight at 4 °C (see Table 2). Note that different lysis and wash buffer recipes were used for experiments intended for western blot or MS proteomics and those intended for RNA sequencing.
5. Chill an aliquot of DPBS to be dosed with CHX (20 mL per 10 cm plate).
6. Prepare the RNAse removal buffer, salt wash, lysis buffer, and wash buffers (see Table 2). For buffers with dense/viscous components such as glycerol and Triton X-100, pipet the completed buffer to ensure even mixing. Chill buffers on ice prior to use.
   NOTE: For bead washing steps, briefly spin the tube containing beads in their current buffer in a pocket centrifuge to collect the contents at the bottom. Place the tube in a magnetic rack for 1 min or until beads have fully migrated. Aspirate the supernatant without touching the bead pellet. Without letting the bead pellet dry, add the next buffer. To resuspend the bead pellet, either pipet gently or roll the closed tube between one’s fingers while holding it against the magnetic rack, such that the beads are quickly pulled in multiple directions by the magnet. Finally, invert the resuspended beads several times to ensure the inside walls of the tube are washed.
7. Resuspend the stock bottle of streptavidin beads by gentle pipetting or rocking. Transfer 450 µL of beads to a protein low-binding tube for each 10 cm plate of mESCs.
8. Following the above NOTE on bead washing, discard the bead storage buffer and wash beads 2x with 900 µL of RNAse removal buffer.
   NOTE: Proceed to the next step without delay; extended time in RNAse removal buffer may decrease the beads’ binding capacity.
9. Wash beads 1x with 900 µL of salt wash.
10. Wash beads 2x with 900 µL of Wash I Buffer. After resuspending beads in the second round of Wash I buffer, store the beads in the Wash I buffer at 4 °C while performing activation and cell lysis.
11. Add biotin (50 µM final concentration; 1:1,000 dilution of 50 mM stock) and CHX (100 µg/mL final concentration; 1:1,000 dilution of 100 mg/mL stock) as appropriate to the warmed aliquot(s) of media. Keep at 37 °C until use.
12. Add CHX (100 µg/mL final concentration; 1:1,000 dilution of 100 mg/mL stock) to the cold aliquot of DPBS and keep chilled until use.
    NOTE: The following two steps describe activation of ALIBi via supplemental biotin and blue light illumination. An ideal duration of activation is long enough to yield an adequate amount of ribosome biotinylation for downstream assays, but short enough to preserve spatiotemporal resolution. Ideal activation time may vary based on parameters such as the cell type, scale of the experiment, expression level of the split enzymes, abundance of ribosomes near the organelle of interest, and sensitivity of the analysis technique. We recommend 15 minutes as a starting point, which is the activation time used in our representative results (Figure 2) and in our accompanying work³⁰.
13. Aspirate media from the transfected plates. Immediately add 10 mL of the warm media containing CHX ± biotin per plate.
14. For any negative control plates not to be illuminated, place them in a dark incubator. For plates to be illuminated, place them directly on top of the transilluminator, turn on the transilluminator, and incubate for 15 min.
15. When illumination is complete, immediately wash the plates 2x with 10 mL of cold DPBS + CHX per plate. Place the plates on ice between and after the washes. After aspirating the second wash, tilt the plate and aspirate any residual DPBS.
16. Keeping each plate as flat as possible on ice, proceed to lysis by adding 2 mL of ice-cold lysis buffer. Incubate plate on ice for 5 min, tapping plate to dislodge cells 2–3x during that time.
    NOTE: if kept dark under foil and on ice, plates can be moved out of the cell culture area to another dark space during this time.
17. Dislodge any remaining cells by using a micropipette to pipet the lysate up and down on the plate surface while avoiding formation of bubbles, then transfer lysate to a chilled and labeled protein low-binding tube.
    NOTE: OPTIONAL: For some difficult-to-lyse organelles such as the nucleus and nucleolus, sonication may help.
18. In the case of difficult-to-lyse organelles, sonicate all samples in the experiment for 45 s at 4 °C (power setting of 3 watts). If the sonicator design requires insertion of a probe into an open sample tube, transfer the lysate initially from the plate into a larger tube to prevent overflow during sonication; then, transfer it to a 2 mL tube.
19. Incubate the lysate on ice for 15 min.
20. Centrifuge the lysate at 17,000 × g for 3 min at 4 °C. Transfer the supernatant (clarified lysate) to a chilled protein low-binding tube without disturbing the pellet of cell debris.
    NOTE: If RNA-seq analysis is intended, an aliquot of the clarified lysate can now be collected as the total RNA fraction (see Supplemental File 1, “Recommendations for RNA sequencing” step 1.1) for details.
    NOTE: If the cell quantities, transfection/culture conditions, and lysis conditions have been uniform for all samples, we typically assume lysate concentrations are comparable between samples and proceed without quantifying total protein concentration. This minimizes the elapsed time between activation and affinity purification.

4. Affinity purification

1. Pellet the washed streptavidin beads using the magnetic rack. Aspirate the Wash I buffer.
2. Transfer 1,200 µL of clarified lysate to the tube containing the washed beads. Pipet gently to resuspend the beads. Store the remainder of the clarified lysate at -80 °C.
3. Incubate the lysate with beads at 4 °C in the dark for 1 h on the tube rotator.
4. During this incubation, prepare the TEV elution buffer (Table 2).

5. Elution

1. Following the notes on bead washing in section 3, pellet the beads on a magnetic rack and transfer the supernatant to a chilled protein low-binding tube. Store the supernatant at -80 °C.
2. Wash beads 2x with 900 µL of Wash I buffer.
3. Wash beads 1x with 900 µL of Wash II buffer.
   NOTE: After completion of this step, samples no longer need to be handled in the dark.
4. Add 450 µL of TEV elution buffer to the beads.
5. Incubate beads in TEV elution buffer for 1 h at room temperature on the tube rotator. For smaller-scale experiments using low volumes of TEV elution buffer, incubate the tubes on a shaker at 1,000–2,000 rpm instead.
6. Pellet the beads on the magnetic rack. Transfer the eluate to a protein low-binding tube. If proteomics analysis or western blot is intended, store the eluate at -80 °C. If RNA-seq analysis is intended, see Supplemental File 1 “Recommendations for RNA sequencing” step 1.2 for eluate storage instructions.
7. OPTIONAL: To preliminarily determine whether the affinity purification has yielded appreciable rRNA/mRNA (indicating that ribosomes have been isolated), analyze eluates on a microvolume spectrophotometer using TEV elution buffer as a blank.
   NOTE: Eluates from cells containing the activated system should have greater absorbance at 260 nm than eluates from cells lacking the split enzymes, blue light/biotin, or L31-FTA.

6. Western blot

NOTE: Until lysate and flow-through samples have been denatured in Laemmli buffer, we recommend continuing to avoid light exposure. We use precast gels, commercially available transfer buffer and running buffer, and a semi-dry transfer system (see the Table of Materials). SDS-PAGE and western blot are performed according to the manufacturers’ instructions for these components, unless otherwise stated below.

1. Add 6x Laemmli buffer to the lysate, flowthrough, and/or eluate samples to a final concentration of 1x Laemmli buffer. Heat at 95 °C in a heat block or thermocycler for 10 min. If condensation forms on the tube lid/walls, spin down tubes briefly.
2. Set up a gel with running buffer in the gel tank, per the tank manufacturer’s instructions. Load the samples and the ladder.
3. Run the gel at 125 V for 45–75 min until the dye front approaches the bottom of the gel. In the meantime, prepare PBST and blocking buffers (Table 2).
4. Perform a semi-dry transfer per the transfer system manufacturer’s instructions.
   NOTE: For all membrane washing/incubation steps, ensure that the membrane is fully submerged and place the membrane tray on an orbital or tilting rocker. For a 10 x 15 cm tray, this requires 25–50 mL of buffer.
5. Remove the membrane from the transfer stack and proceed to the next step immediately to ensure the membrane does not dry out.
   NOTE: OPTIONAL: Dry spots on the membrane can result in uneven blocking or probe binding, especially with probes that have short incubations such as streptavidin-horseradish peroxidase. To avoid this issue, we re-wet membranes after transfer in 100% methanol for 30 s, then in water for 5 min. This measure is similar to general recommendations for re-wetting PVDF membranes that have dried after transfer^32,33.
6. Wash the membrane in PBST for 3 x 10 min at room temperature.
7. Following the specified conditions in Table 3 for each western blot probe, perform the blocking, washing, probe incubation, and secondary antibody incubations.
8. After the final wash in step 6.7, mix the ECL substrate components per the manufacturer’s instructions and coat the membrane evenly with it. Capture the image on a chemiluminescence gel imaging system.

结果

Western blot（方案第6部分）是初步评估实验是否成功最便捷的方法。当使用链霉亲和素-HRP探针检测膜时，所有系统组分均存在的细胞裂解液中应出现一条对应于生物素标记的L31-FTA的条带（图2A，第20泳道）。如果细胞转染了全长BirA作为阳性对照，则同一条带通常会更明显（图2A，第1–4泳道）。当缺失一个或多个系统组分时，该条带应变弱或消失（图2A，其余所有泳道）。由于哺乳动物细胞内源性含有多种分子量在70–80 kDa和120–130 kDa范围内的生物素化羧化酶蛋白^34,35，因此在所有裂解液样本中通常均可观察到这些分子量的条带。此外，也可使用抗V5抗体探针膜以确认分裂BirA片段的表达。

通过蛋白质印迹（Western blot）检测裂解液、穿流液和洗脱液样品，可以评估亲和...

讨论

ALIBi offers several advantages relative to existing techniques for studying subcellular translation and has potential future applications in ribosome biology and beyond. This protocol requires similar or less time compared to sequential lysis and centrifugation^21,22 and is potentially applicable to any subcellular compartment that can be genetically targeted with a localizing peptide sequence. Compared to constitutively active BirA^10...

材料

本文使用的材料清单

姓名	公司	目录编号	评论
-80 °C存储			用于细胞培养
0.1%明胶水溶液	MilliporeSigma	ES-006-B	用于细胞培养
0.2 µm滤器单元	Fisher	04-977-158	用于细胞培养
0.5%胰蛋白酶-EDTA (10X)	ThermoFisher	15400-054	用于细胞培养
1 M MgCl2	ThermoFisher	AM9010	用于裂解和亲和纯化
1 M Tris pH 7.5	ThermoFisher	15567027	用于裂解和亲和纯化
1 M Tris pH 8	ThermoFisher	AM9855G	可选：用于将FTA插入内源性核糖体蛋白基因座
1.5和2 mL微离心管，经高压灭菌			用于转染
1.5、2 mL和5 mL低结合蛋白微离心管	Eppendorf	022431081, 022431102, 0030108302	用于裂解和亲和纯化
10 cm组织培养皿			用于细胞培养
10 M NaOH	MilliporeSigma	72068-100ML	用于裂解和亲和纯化
10x Tris/甘氨酸/SDS 运行缓冲液浓缩液	Bio-Rad	1610772	用于蛋白质印迹
37 °C组织培养箱 x 2			用于细胞培养。其中一个作为细胞培养的主培养箱和转染细胞的照明装置。在激活ALIBi的当天，在第二个培养箱中需要临时空间以容纳转染的细胞，同时在主培养箱中设置荧光照明器。
37 °C水浴			用于细胞培养
5 M NaCl	ThermoFisher	AM9010	用于裂解和亲和纯化
50 mM生物素	ThermoFisher	B20656	用于裂解和亲和纯化
6x Laemmli缓冲液	FIsher	50-196-784	用于蛋白质印迹
70%乙醇			用于消毒
乙腈	Fisher	A955	可选：用于质谱样品制备
AcTEV蛋白酶试剂盒	ThermoFisher	12575015	用于裂解和亲和纯化
ALIBi C-片段质粒	Addgene	#235615, 235636 – 235641	转染需要Midi-prep规模
ALIBi N-片段质粒	Addgene	#235614, 235635, 235642, 235643	转染需要Midi-prep规模
铝箔			用于裂解和亲和纯化
碳酸氢铵	MilliporeSigma	5.3305	可选：用于质谱样品制备
抗FLAG抗体	MilliporeSigma	F3165	用于蛋白质印迹
抗GAPDH抗体	Cell Signaling	5174	用于蛋白质印迹
抗小鼠二抗	GE Healthcare	NA931V	用于蛋白质印迹
抗兔二抗	GE Healthcare	NA934V	用于蛋白质印迹
抗Rpl12抗体	Abcam	ab175219	用于蛋白质印迹
抗Rpl31抗体	Abcam	ab103991	用于蛋白质印迹
抗Rpl5抗体	ProteinTech	15430-1-AP	用于蛋白质印迹
抗V5抗体	ThermoFisher	R960-25	用于蛋白质印迹
磷酸二酯酶	NEB	M0398L	用于裂解和亲和纯化
β-巯基乙醇（55 mM）	ThermoFisher	21985-023	用于细胞培养
蓝光荧光照明器	Clare Research Chemical	DR46B（小）或 DR196（大）	用于裂解和亲和纯化
牛血清白蛋白（BSA）	Fisher	BP1600-100	用于蛋白质印迹
C18提取尖端（OMIX C18提取尖端）	Agilent	A57003100	可选：用于质谱样品制备
细胞冷冻容器			用于细胞培养
适用于15 mL和/或50 mL锥形管的离心机			用于细胞培养
适用于1.5-2 mL微离心管的离心机			用于裂解和亲和纯化。应该能够冷藏至4 C，并且速度可达17,000 g。理想情况下，它位于一个可以在转移管子进出时暂时变暗的房间里。
Clarity ECL试剂盒	Bio-Rad	170-5061	用于蛋白质印迹
Criterion凝胶槽	Bio-Rad	1656001	用于蛋白质印迹
Criterion TGX预成型凝胶，4-20%	Bio-Rad	5671095	用于蛋白质印迹。还可提供其他孔径和布局
环己酰亚胺	MilliporeSigma	C7698-5