Auxin-inducible degron system for depletion of proteins in S. cerevisiae

Abstract

The auxin-inducible degron (AID) is a powerful tool that is used for depletion of proteins to study their function in vivo. This method can conditionally induce the degradation of any protein by the proteasome, simply by the addition of the plant hormone auxin. This approach is particularly valuable to study the function of essential proteins. The protocols provided here describe the steps to construct the necessary strains and to optimize auxin-inducible depletion in S. cerevisiae.

BASIC PROTOCOL 1:

CONSTRUCTION OF TIR1-EXPRESSING STRAINS BY TRANSFORMATION

BASIC PROTOCOL 2:

TAGGING A YEAST PROTEIN OF INTEREST WITH AN AUXIN-INDUCIBLE DEGRON

SUPPORT PROTOCOL 1:

CONSTRUCTION OF DEPLETION STRAINS BY GENETIC CROSSES

BASIC PROTOCOL 3:

OPTIMIZATION FOR DEPLETION OF THE AID-TAGGED PROTEIN

INTRODUCTION

One of the most effective methods to study gene function in vivo is to reduce or eliminate the gene product, which can be achieved by mutation, transcriptional repression of the gene, or destabilization of the gene product. In S. cerevisiae, for non-essential genes, this is easily accomplished by gene deletions (Giaever et al., 2002), although the results can be complicated by secondary effects (Hughes et al., 2000; Kwan, Wang, Amemiya, Brewer, & Raghuraman, 2016; Teng et al., 2013). For essential genes, different conditional approaches have been used to deplete the gene product, including temperature-sensitive mutations (Costanzo et al., 2016), mechanisms to reduce transcript levels (Breslow et al., 2008; Mnaimneh et al., 2004), and a system to sequester nuclear proteins in the cytoplasm (anchor-away) (Haruki, Nishikawa, & Laemmli, 2008). In addition to these approaches, methods to induce protein degradation have been developed and used, although with mixed success (Dohmen, Wu, & Varshavsky, 1994; Labib, Tercero, & Diffley, 2000; Moqtaderi, Bai, Poon, Weil, & Struhl, 1996).

More recently, a highly efficient method for inducible protein degradation has been developed. This approach relies on the evolutionarily conserved SCF degradation pathway and uses additional target-specific components derived from plants. In plants, the hormone auxin mediates the interaction of the protein TIR1 (Transport Inhibitor Response 1 protein) with the AUX/IAA family of transcriptional repressors, leading to AUX/IAA degradation. TIR1 is part of the E3 ubiquitin ligase SCF-TIR1 that recruits an E2 ubiquitin-conjugating enzyme. Together, they polyubiquitinate and target AUX/IAA proteins via a specific amino acid degron sequence for proteasome-dependent degradation. To adapt this system to yeast for targeted degradation of any protein of interest, investigators expressed TIR1 from Oryza sativa (OsTIR1) in yeast and fused the Arabidopsis thaliana IAA7 auxin-inducible-degron (AID) to the protein of interest (Nishimura, Fukagawa, Takisawa, Kakimoto, & Kanemaki, 2009). As the other members of the ubiquitin-dependent pathway are conserved, these yeast strains can be treated with auxin to induce proteasomal degradation of the AID-tagged protein (Figure 1).

Figure 1.

Depletion of protein by the AID system. The protein of interest (POI) is fused to an AID tag in a yeast strain that expresses the TIR1 protein. Addition of auxin promotes the interaction between TIR1-containing Skp1-F-box-Cullin E3 ubiquitin ligase, which recruits the E2 ubiquitin conjugating enzyme. Together they polyubiquitinate the AID tag, thereby targeting the AID-tagged protein for degradation by the proteasome. Image created with BioRender.

This method has been successfully used to study loss of function for essential proteins in a variety of biological processes (for example, (Eng, Guacci, & Koshland, 2014; Snyder et al., 2019)). The depletion of the degron-tagged protein is rapid and efficient, such that within 30–60 minutes the protein of interest is depleted to less than 10% that of wild-type levels. There is minimal strain construction involved, and the method is cost-effective and feasible for genome-wide experiments.

The degron approach to deplete a specific protein requires two components within the same yeast strain: (1) constitutive expression of the OsTIR1 protein and (2) the degron tag fused to the target protein. The steps described below provide details to construct the yeast strains and to optimize conditions for protein depletion. Another review has also described the steps in this protocol (see Current Protocols article: Nishimura & Kanemaki, 2014).

BASIC PROTOCOL 1

CONSTRUCTION OF TIR1-EXPRESSING STRAINS BY TRANSFORMATION

A plasmid containing the OsTIR1 gene with targeting sequences upstream and downstream of the LEU2 locus as previously described (Chan, Mugler, Heinrich, Vallotton, & Weis, 2018) and depicted in Figure 2 can be used to generate a strain expressing the OsTIR1 protein under the control of the S. cerevisiae GPD1 promoter using the following steps.

Figure 2.

pTIR1 plasmid and restriction fragment for transformation of yeast. The pTIR1 plasmid (9719 bp) has regions homologous (gray) to 646 bp upstream and 525 bp downstream of the S. cerevisiae LEU2 gene. Between these homology regions is the Candida glabrata LEU2 (Cg LEU2) gene with its own promoter and terminator, and the OsTIR1 gene is expressed under the control of the S. cerevisiae GPD1 promoter. The Cg LEU2 gene complements the S. cerevisiae leu2Δ mutation, thus allowing selection of yeast containing the plasmid fragment on medium lacking leucine. The fragment (6221 bp) used for transformation is released by digestion with Pme1. The arrows indicate the direction of genes. Os: Oryza sativae, Ca tADH1: Candida albicans ADH1 terminator. Image created with BioRender.

Materials

E. coli strain with the plasmid pTIR1 (pKW2830 in ((Chan et al., 2018); available upon request from the authors of this article). The GPD1-TIR1 construct expresses TIR1 constitutively when glucose is used as a carbon source in the growth medium.

Yeast strain such as MATa leu2Δ1. (A leu2Δ mutation is required to select for integration of the pTIR1 plasmid, pKW2830.)

PmeI restriction enzyme

Plasmid miniprep kit and DNA purification kit from agarose gels, such as the E.Z.N.A. kits from Omega Bio-tek

Primers to check integration at the S. cerevisiae LEU2 locus by colony PCR. Primers can be designed using a strategy similar to the one described in Basic Protocol 2 (Figure 4)

Figure 4.

Checking the AID tag by colony PCR. A PCR reaction with three primers is set up. Primers ‘a’ and ‘b’ will give two different product sizes for the two possible different alleles (with and without the AID sequence), whereas primers ‘a’ and ‘c’ will generate a product only for the correct fusion. GOI: gene of interest. Image created with BioRender.

SC-Leu dropout plates (see recipe)

1. Digest the pTIR1 plasmid with homology regions to target locus as described below.

   The pTIR1 plasmid is used to generate a fragment that contains the OsTIR1 gene flanked by sequences homologous to ~500 bp upstream and downstream of the S. cerevisiae LEU2 gene (Figure 2).

   1. Perform restriction digestion of 10–20 μg of the pTIR1 plasmid using PmeI.
      This results in a linear fragment containing the Candida glabrata LEU2 gene and the OsTIR1 gene flanked by sequences homologous to DNA sequences upstream and downstream of the S. cerevisiae LEU2 gene.
   2. Gel-purify the digested fragment (6221 bp) using a gel purification kit or protocols for agarose gel purification of DNA (see Current Protocols article: Moore, Dowhan, Chory, & Ribaudo, 2002) and resuspend the DNA in approximately 10 μL, for transformation of S. cerevisiae.

2. Transform the desired yeast strain (see Current Protocols article: Becker & Lundblad, 2001) with the purified restriction fragment and select for transformants on an SC-Leu dropout plate.

3. Single-colony purify the Leu⁺ transformants on SC-Leu plates and then test by colony PCR for the presence of the OsTIR1 construct. This can be done using a DNA polymerase such as the KAPA2G Robust HotStart polymerase. The PCR check can be done with primers that anneal to S. cerevisiae LEU2 sequences that flank the integrated construct. Different sized products are obtained before and after integration. Always include the parental strain as a control. The colony PCR is setup as follows: a small amount of the colony is suspended in 50 μL 20 mM NaOH and incubated for 5–10 minutes at 95°C. 2.5 μL of the boiled mix is used in a 25 μL PCR reaction containing: Buffer B (1x), Enhancer (1x), dNTPs (0.2 mM each), three primers (0.5 μM each), and KAPA2G polymerase (0.4 U).

   The TIR1-expressing strain thus generated can then be used for tagging the gene of interest with an AID tag for depletion by auxin.

BASIC PROTOCOL 2

TAGGING A YEAST PROTEIN OF INTEREST WITH AN AUXIN-INDUCIBLE DEGRON

Auxin-dependent conditional depletion of a protein requires the protein to be fused to an IAA7 degron tag (hereafter referred to as an AID tag) in a strain that expresses the gene encoding the OsTIR1 protein. Below is the detailed protocol to construct this fusion, starting with a yeast strain such as was constructed in Basic Protocol 1. In this protocol, a DNA cassette encoding the AID tag is generated by PCR, followed by fusion to the gene of interest by transformation and homologous recombination.

Materials

E. coli strain with plasmid pScAID2 (Eng et al., 2014)

Plasmid miniprep kit and DNA purification kit such as the E.Z.N.A. kits from Omega Bio-tek

LB plates and LB liquid medium with ampicillin added to a concentration of 50 μg/mL

Restriction enzymes to check the plasmid by digestion

High-fidelity polymerase for PCR such as KAPA2G Robust HotStart DNA polymerase

Yeast strain expressing the OsTIR1 gene (Basic Protocol 1)

YPD plates (see recipe)

YPD+G418 plates (see recipe)

Anti-V5 antibody such as that commercially available from Invitrogen (RRID: AB_2556564)

Tubes for growing 5 mL yeast cultures

30°C incubator

Spectrophotometer

Hemocytometer and phase-contrast microscope

Roller drum and shaker for growing cultures

Reagents required for yeast transformation (see Current Protocols article: Becker & Lundblad, 2001) and western blotting (see Current Protocols articles: Matsuo, Asakawa, Toda, & Katayama, 2006; Ni, Xu, & Gallagher, 2016)

Plasmid preparation:

The template plasmid pScAID2 (Figure 3), containing the AID tag along with the KanMX6 resistance cassette, is prepared using a plasmid miniprep kit. The plasmid is then used as a template for PCR to generate a fragment to be used for transformation of the yeast strain expressing the OsTIR1 protein.

Figure 3.

pScAID2 plasmid. The cassette carrying 3xV5-IAA7-KanMX6 is amplified with two primers as indicated. The 3’ part of the primers has 40 base pairs of homologous sequence to the site where the tag is to be integrated. GOI: gene of interest. Image created with BioRender.

• 1Inoculate a single colony of E. coli that contains plasmid pScAID2 into 5 mL LB containing ampicillin and grow overnight at 37°C with shaking.
• 2Perform a miniprep using a kit such as the E.Z.N.A. plasmid mini kit. Typically, one obtains about 25 μg of plasmid from each 5 mL culture, which is sufficient for use as a template for PCR.
• 3It is advisable to check the plasmid by restriction digestion to ensure the correct structure. For example, a diagnostic digestion with HindIII, which cuts the plasmid three times, should result in linear fragments of 2955 bp, 1585 bp and 492 bp. Furthermore, the size of the plasmid (5032 bp) can be confirmed by linearizing the plasmid using an enzyme such as EcoRV, which digests the plasmid once.

Primer design to amplify the DNA sequence encoding the AID tag:

The pScAID2 plasmid is constructed in the pFA6a backbone (Eng et al., 2014; Longtine et al., 1998). The plasmid has a 3xV5 sequence encoding a V5 epitope tag in frame with the IAA7 sequence that has been codon-optimized for yeast, followed by the KanMX6 resistance cassette. Primers are designed such that the amplified 3xV5-AID-KanMX6 DNA sequence from the pScAID2 plasmid will be flanked on either side by 40 bp of homologous sequences to the 3’ end of the target gene (Figure 3).

• 4The forward primer is designed by choosing 40 nt of the coding strand (sense strand) of the gene immediately upstream from the stop codon of the target gene to be tagged. The stop codon is excluded from this primer. This is followed by the 20 nt of the forward primer sequence to amplify the 3xV5-AID-KanMX6 sequence from the pScAID2 plasmid. Thus, the 60 nt forward primer will be: 5’- (40 bp of sense sequence of the target gene)- CGGATCCCCGGGTTAATTAA −3’.
• 5The reverse primer is designed by choosing 40 nt of the non-coding strand (antisense strand) of DNA immediately downstream from the stop codon of the target gene to be tagged. This is followed by the 20 nt of the reverse primer sequence to amplify the 3xV5-AID-KanMX6 sequence from the pScAID2 plasmid. Thus the 60 nt reverse primer will be: 5’- (40 bp of antisense sequence downstream of the target gene)- GAATTCGAGCTCGTTTAAAC −3’.
• 6The primers are synthesized at 25 nmol scale, purified using standard desalting, and resuspended in PCR grade water to a stock concentration of 100 μM.

PCR amplification to generate DNA for transformation of yeast

• 7PCR amplify the DNA fragment using a high-fidelity polymerase such as Phusion or Q5 polymerase. Perform 30–35 cycles of amplification using the prescribed amplification time and temperature for the enzyme used. Typically, a 250 μL PCR reaction using 50–100 ng of plasmid template generates enough material (approximately 1–10 μg of DNA fragment) for one transformation. Sometimes multiple PCR reactions are performed and then pooled to obtain a greater amount of DNA.
• 8Check the size and amount of the PCR product by electrophoresis on a 0.6% agarose gel. The expected size of the PCR product is 2654 bp (40 bp upstream target sequence + 2574 bp 3xV5-AID-KanMX6 + 40 bp downstream target sequence). The size and amount are judged by comparing to a sample of a known amount of DNA size markers.
• 9DNA can be purified by using a PCR purification kit and then used for transformation. However, to avoid transformation of any non-specific products obtained during PCR, it is recommended to gel-purify the DNA using a kit such as the E.Z.N.A. gel extraction kit.
• 10Quantify the DNA using a nanodrop. Approximately 10 μg of DNA is used per transformation.

Transformation of OsTIR1-expressing yeast strains to tag the target protein with the AID tag

• 11Inoculate a single colony of the yeast strain with TIR1 integrated at the LEU2 locus into 5 mL of YPD and grow to saturation overnight at 30°C.
• 12The next morning, count the cells using a hemocytometer and use the culture to inoculate 10 mL of YPD to an initial concentration of 2.5 × 10⁶ cells/mL. Grow cells at 30°C with shaking 3–5 hours to a concentration of 2 × 10⁷ cells/mL.
• 13Transform the yeast cells using the LiOAc transformation protocol (Gietz & Schiestl, 2007) and spread the transformed yeast onto a YPD plate. A “no DNA control” transformation should always be included.
• 14The next day, replica plate the lawn of yeast from the YPD plate onto a YPD+G418 plate and incubate those plates for 3–4 days at 30°C.
• 15The transformants that appear on YPD+G418 plates are single-colony purified on YPD+G418 plates and incubated for 2 days at 30°C.
• 16G418-resistant colonies are then tested by colony PCR for the presence of the AID tag. This can be done using a DNA polymerase such as the KAPA2G Robust HotStart polymerase. The PCR check can be done with three primers such that different sized products are obtained with either the untagged or the tagged allele for the gene of interest (Figure 4). Always include the parental untagged strain as a control. The colony PCR is setup as follows: a small amount of the colony is suspended in 50 μL 20 mM NaOH and incubated for 5–10 minutes at 95°C. 2.5 μL of the boiled mix is used in a 25 μL PCR reaction containing: Buffer B (1x), Enhancer (1x), dNTPs (0.2 mM each), three primers (0.5 μM), and KAPA2G polymerase (0.4 U).
• 17Colonies that are positive for the AID tag by PCR are then checked further for expression of the AID-tagged protein by western blot.

Western blot analysis of PCR-positive strains to identify strains that express the AID-tagged protein

• 18Inoculate a single colony of the PCR-positive strains to be analyzed into 5 mL of YPD and grow overnight at 30°C. The next morning, dilute cultures into 10 mL of YPD to a density of ~ 3 × 10⁶ cells/mL (OD₆₀₀ ~0.1) and grow at 30°C for 4–6 hours to a density of ~10⁷ cells/mL (OD₆₀₀ ~0.1 ~0.4–0.5). Cultures are pelleted, and the pellets can be stored at −70°C or processed further for western blot analysis.
• 19Protein extraction and western blotting are performed as described (Matsuo et al., 2006; Ni et al., 2016), and the blots are probed with anti-V5 antibody to confirm expression of the proteins with degron tags.

  If antibodies against the native protein are available, then it is advisable to compare the expression of the tagged protein to the native protein levels in untagged strains to characterize the possible effect of the tag on protein levels.

• 20Strains that are confirmed by colony PCR and western blot can then be stored in 15% glycerol at −70°C for permanent storage. Typically, two independent isolates for each strain should be stored.

  It is crucial to check for all genetic markers to ensure that the strain genotype is correct. Typically, a plate containing the tagged strains and an untagged control strain is replica plated onto plates to test drug resistances, nutritional auxotrophies, and mating type.

SUPPORT PROTOCOL 1

CONSTRUCTION OF DEPLETION STRAINS BY GENETIC CROSSES

The strain to be used for depletion studies, expressing both OsTIR1 and the AID-tagged protein of interest, can also be constructed by crosses. Crosses can be done as described in (Current Protocols article:Treco & Winston, 2008) using a strain expressing TIR1 (for example, MATa leu2Δ1::CgLEU2::OsTIR1) and a strain expressing the AID-tagged protein of interest (POI) (MATα ura3–52 leu2Δ1 POI-V5-AID::KanMX6).

Mate the strains by mixing together a purified colony from each on a YPD plate using sterile toothpicks and incubate at 30°C. After 4–5 hours of incubation, check for zygotes. When zygotes are present, separate zygotes on a YPD plate using a micromanipulator and grow them at 30°C for 2 days to obtain diploid colonies. Inoculate a YPD culture with part of a new colony and grow overnight. The following day, dilute the culture to OD₆₀₀ ~ 0.05–0.1 and grow for about three generations (to mid log). Spin down ~ 200 μL of the culture, wash the cells 2–3 times with sterile water, resuspend them in 2 mL of liquid sporulation medium, and grow 3–4 days at 25°C. Dissect tetrads as described (see Current Protocols article:Treco & Winston, 2008). Score tetrads for all genetic markers and store 2–3 strains with the desired genotype as glycerol stocks. These strains are now ready to be analyzed for depletion of the degron tagged protein.

BASIC PROTOCOL 3

OPTIMIZATION FOR DEPLETION OF THE AID-TAGGED PROTEIN

Auxin-dependent conditional depletion is achieved by the addition of auxin to an exponentially growing culture of the yeast strain of interest. The time of treatment required for optimal depletion is empirically determined by performing a time course experiment, measuring protein levels from each time point by western blot. The type of auxin and the auxin concentration to be used can be similarly determined and depend on experimental conditions to be used (see critical parameters).

Once depletion conditions are optimized, one can assay the depleted cells by any method of interest. These might include measurements of gene expression (for example, RNA-seq, ChIP-seq, or ribosome profiling), assay of cell cycle, protein localization, or the effects of second enhancer or suppressor mutations.

Materials

Yeast strain expressing OsTIR1 and AID-tagged protein of interest (Support Protocol 1)

YPD medium (see recipe)

Dropout media to check genetic markers (see recipe)

Auxin, either the natural form (indole-3-acetic acid, IAA) or a synthetic version (naphthalene-acetic acid, NAA). These are commercially available from SIGMA.

DMSO

Anti-V5 antibody

Tubes for growing 5 mL liquid yeast cultures

30°C incubator

Spectrophotometer

Roller drum and shaker for growing cultures

Reagents for ChIP (see Current Protocols article: Aparicio, Geisberg, & Struhl, 2004)(if necessary)

Growth of culture

• 1Inoculate a single colony of the AID-tagged strain into 5 mL of liquid YPD in a test tube.
• 2Incubate overnight at 30°C on a roller drum or with shaking to obtain a saturated culture. This will be the starter culture to be used for inoculating a larger culture the next day.
• 3The next day calculate the volume of the starter culture needed to inoculate 130 mL of liquid YPD medium using the formula below. Incubate at 30°C with shaking. The objective is to obtain a culture with a final density of ~10⁷ cells/mL (OD₆₀₀ ~ 0.4–0.5) at the desired time.

  Formula for calculating volume of the starter culture to be used (Sabatinos & Forsburg, 2010): SV = (TV * OD_final) / (OD_start * 2 ^(t/g−1)). SV= volume of starter culture to be used, TV = total volume, OD_final = final O.D. desired, OD_start = O.D. of starter culture, t= time of incubation in hours and g= doubling time of the culture in hours (typically 1.5 hours for wild-type yeast grown in YPD).

• 4When the OD₆₀₀ is ~0.4–0.5, divide the culture into 5 flasks of 25 mL each. These cultures can now be used to perform a time-course experiment to determine the optimal auxin treatment time for depletion of the AID-tagged protein. Before proceeding to the next step, spot a drop of the culture onto a YPD plate and streak for single colonies followed by replica plating to check for growth phenotypes on dropout plates to ensure that the culture contains the correct strain without any contaminants.

Time course experiment to determine the optimal time for depletion

• 5Prepare a stock of the auxin IAA (or NAA, see note under Critical Parameters) in DMSO. Typically, a fresh stock of 50 mM IAA is prepared.
• 6Label the five flasks ‘a’ through ‘e.’ Then add DMSO to flask ‘a’ and auxin to a final concentration of 100 μM to flasks b-e.
• 7Incubate all flasks at 30°C with shaking.
• 8Remove flasks from the incubator at the desired time points and pellet 20 mL of the culture by spinning at 2500 × g for 5 minutes in a bench-top centrifuge. Use the rest of the culture for measuring viability and assaying phenotypes as described below. The times chosen for the depletion time course will depend on the specific AID-tagged protein. Initially, a time course of 30, 60, 90, and 180 minutes can be used. Process the control culture with DMSO (flask ‘a’) along with the sample for the last time point.
• 9Store the cell pellets at −80°C or process immediately for analysis by western blot.

Measuring AID-tagged protein depletion by western blot

• 10Prepare protein extracts from the stored pellets as described in (Matsuo et al., 2006). Perform western blotting using anti-V5 antibody, and determine the optimal treatment time and concentration for depletion of proteins based on these results (Figure 5). The optimal conditions (see note under “Analyzing depletion” in Critical Parameters) can then be used in future experiments.

Figure 5.

Time course experiment to determine the time required for depletion of the protein of interest. The Spn1–3xV5-AID tagged strain was grown in YPD and treated with 100 μM IAA for the indicated time (in minutes), and protein extracts were prepared. The levels of Spn1 protein were analyzed by western blotting using anti-Spn1 antisera. Pgk1 protein was used as a loading control.

Measuring cell viability after the depletion of essential AID-tagged proteins

• 11To measure the total number of cells, pellet 1 mL of culture at each time point, including the DMSO control culture, wash the cells with 1 mL of water twice by resuspension and centrifugation, and then finally resuspend the pellet in 1 mL of water. Count the cells using a hemocytometer.
• 12To measure the number of viable cells, make serial dilutions of each culture and plate them onto YPD to ideally obtain 100–150 colonies per plate. Incubate plates for two days at 30°C, and then perform a colony count to determine the percent viability for each culture compared to the DMSO control culture.

REAGENTS AND SOLUTIONS

YPD

Prepare the following in Flask 1 (1-L):

Yeast extract 10 g

Peptone 20 g

dH₂O 500 mL

Prepare the following in Flask 2 (2-L):

Agar 20 g

dH₂O 440 mL

Autoclave the two flasks, then mix the contents together in the 1-L flask. Add 50 mL of sterile 40% glucose (sterilized by autoclaving separately) and 10 mL of 40 mM tryptophan (filter-sterilized and stored at 4°C). Allow the media to cool a bit before pouring into Petri plates. Note: 1 L of media should yield ~35–40 100-mm plates.

YPD + G418–200 μg/mL

Follow YPD recipe above. Dissolve G418 in 1–2 mL of dH₂O. Filter-sterilize with a syringe filter. Add to media as it is cooling. Note: The amount of G418 powder depends on the activity of the specific lot.

Dropout Plates: (e.g. SC-Leu)

Prepare the following in Flask 1 (1-L):

Dropout mix (available from vendors such as Takara Bio, described in (Sherman, Fink, & Hicks, 1987), and can be found at many web sites) 2 g

Yeast nitrogen base (w/o ammonium sulfate and amino acids) 1.45 g

Ammonium sulfate 5 g

dH₂O 500 mL

Prepare the following in Flask 2 (2-L):

Agar 20 g

dH₂O 450 mL

Autoclave the two flasks, then mix the contents together in the 1-L flask. Add

50 mL of sterile 40% glucose. Allow the media to cool a bit before pouring into Petri plates.

COMMENTARY

Background Information

Conditional depletion of a protein is a powerful tool for studying the function of a protein in yeast. There are several important advantages to auxin-inducible depletion. First, it allows the study of essential proteins Second, this method is fast and efficient. Typically, most proteins are depleted to below 10% within 30–60 minutes after the addition of auxin. This is valuable for studying the immediate consequences of loss of protein function. Third, auxin causes relatively mild effects on yeast metabolism and gene expression (Nishimura et al., 2009; Prusty, Grisafi, & Fink, 2004). Fourth, the degron tag described here uses a codon optimized version of the AID tag and an OsTIR1 gene expressed under the control of the promoter of the yeast GPD1 gene, which ensures reliable and constitutive expression of TIR1 protein (Chan et al., 2018; Eng et al., 2014). Finally, the method is relatively simple and can be adapted to a variety of genomic and proteomic approaches for studying protein function (for example, (Doris et al., 2018; Gopalakrishnan, Marr, Kingston, & Winston, 2019; Shetty et al., 2017)).

Critical Parameters

Strain construction

Care should be taken to ensure integrity of the plasmids, restriction fragments, and PCR products used for transformation. It is advisable to gel purify the fragments used for transformation and sequence the tagged locus to ensure the integrity of the protein of interest. Also, it is imperative to confirm the protein-tagging by western blot after confirming the integration of the tag by PCR to ensure appropriate expression before proceeding with the depletion experiment.

Choice of auxin

The depletion of degron-tagged proteins can be achieved using either the natural form of auxin, IAA (indole-3-acetic acid), or a synthetic version, NAA (naphthalene-acetic acid). We have found that both IAA and NAA can be reliably used to deplete proteins in liquid culture (Doris et al., 2018; Gopalakrishnan et al., 2019; Shetty et al., 2017) as well as on plates. However, certain conditions such as fluorescent light can affect the stability of the auxin and thereby necessitate the use of NAA over IAA (Papagiannakis, de Jonge, Zhang, & Heinemann, 2017). It is advisable to perform a depletion time course and growth on liquid and solid media using both forms of auxin to determine the best source for the particular protein of interest and experimental conditions used.

The optimal concentration of auxin to be used for depletion of the AID-tagged protein should be determined empirically, as different yeast strains might have different sensitivities to auxin. In general, one wants to use the lowest concentration of auxin that is sufficient for efficient depletion in order to avoid any undesirable effects due to the addition of auxin. The protocol described above uses 100 μM IAA; however, we have found that concentrations as low as 25 μM IAA are sufficient to achieve efficient depletion. One can test a range of concentrations of auxin, for example from 10 μM - 500 μM, and determine the minimal concentration required for effective depletion of the AID-tagged protein. A concentration curve can be performed in similar fashion to the one described for determining the optimal time of depletion.

Analyzing depletion

Depletion should be quantified using a variety of methods. Western blotting analyzes the bulk level of protein in a population of cells. For chromatin-associated proteins, ChIP should be performed to ensure depletion of the protein from chromatin. Additionally, phenotypic assays and microscopy (if used with fluorescent-tagged AID) (Papagiannakis et al., 2017) can be performed to get a better estimate of the extent of depletion.

For depletion of essential proteins, it is crucial to test the viability of the strains in the time course experiment to ensure that one is working with live cells at the time of harvesting. Viability can be tested using the method described above, by micromanipulation of single cells (Shetty et al., 2017), or by vital staining.

The optimal time for depletion will depend on the AID-tagged protein and the purpose of the experiment. For instance, in the case of essential proteins the optimal time of depletion is generally the earliest time at which the AID-tagged protein is depleted without causing loss of viability. However, other considerations, such as the time required for biological effects of depletion of the AID-tagged protein to be measurable, could require adjusting the optimal time.

Troubleshooting

PCR and strain constructions

For construction of yeast strains by transformation with PCR-generated DNA fragments, KAPA2G robust HotStart generally provides sufficient fidelity. For a higher-fidelity polymerase, one can use other enzymes, such as Phusion or Q5 polymerase. After identification of transformants, generally over 50% of the candidates have the desired integration. If the transformation efficiency is too low, increasing the homology for targeting or the amount of DNA to be transformed might be helpful.

Protein depletion

In certain cases, if depletion using the described strategy is not efficient, it can be combined with other methods, such as transcriptional repression, as described for fission yeast (Kanke et al., 2011; Shetty et al., 2017) and for budding yeast (Moqtaderi et al., 1996; Tanaka, Miyazawa-Onami, Iida, & Araki, 2015). The AID system might be sensitive to higher temperatures (Nishimura et al., 2009), and this should be taken into consideration if depletion assays are combined with experimental conditions that require growth at higher temperatures, although the OsTIR1 system used here is stable over a range of temperatures (Nishimura et al., 2009).

Understanding the Results

The size of the PCR product generated for AID-tagging, to be used for transformation of yeast in Basic Protocol 1, yields a 2654-bp product. This will change if longer homology regions are used. Depletion of most proteins is achieved within 30–60 minutes after addition of auxin.

Time Considerations

Construction of the yeast strain with TIR1 integrated at the LEU2 locus takes about one week. Construction of the protein-AID fusion takes about two weeks, starting from PCR amplification of the AID tag through confirmation of depletion by western blotting. To save time, the AID-tagged and TIR1 strains can be generated simultaneously in different strains of opposite mating type and the final strain can be constructed by a cross (Support Protocol 1). This will take about two weeks and the time required for depletion can then be determined in about four days.