Growth-based Determination and Biochemical Confirmation of Genetic Requirements for Protein Degradation in Saccharomyces cerevisiae

Summary

This article describes a yeast growth-based assay for the determination of genetic requirements for protein degradation. It also demonstrates a method for rapid extraction of yeast proteins, suitable for western blotting to biochemically confirm degradation requirements. These techniques can be adapted to monitor degradation of a variety of proteins.

Abstract

Regulated protein degradation is crucial for virtually every cellular function. Much of what is known about the molecular mechanisms and genetic requirements for eukaryotic protein degradation was initially established in Saccharomyces cerevisiae. Classical analyses of protein degradation have relied on biochemical pulse-chase and cycloheximide-chase methodologies. While these techniques provide sensitive means for observing protein degradation, they are laborious, time-consuming, and low-throughput. These approaches are not amenable to rapid or large-scale screening for mutations that prevent protein degradation. Here, a yeast growth-based assay for the facile identification of genetic requirements for protein degradation is described. In this assay, a reporter enzyme required for growth under specific selective conditions is fused to an unstable protein. Cells lacking the endogenous reporter enzyme but expressing the fusion protein can grow under selective conditions only when the fusion protein is stabilized (i.e. when protein degradation is compromised). In the growth assay described here, serial dilutions of wild-type and mutant yeast cells harboring a plasmid encoding a fusion protein are spotted onto selective and non-selective medium. Growth under selective conditions is consistent with degradation impairment by a given mutation. Increased protein abundance should be biochemically confirmed. A method for the rapid extraction of yeast proteins in a form suitable for electrophoresis and western blotting is also demonstrated. A growth-based readout for protein stability, combined with a simple protocol for protein extraction for biochemical analysis, facilitates rapid identification of genetic requirements for protein degradation. These techniques can be adapted to monitor degradation of a variety of short-lived proteins. In the example presented, the His3 enzyme, which is required for histidine biosynthesis, was fused to Deg1-Sec62. Deg1-Sec62 is targeted for degradation after it aberrantly engages the endoplasmic reticulum translocon. Cells harboring Deg1-Sec62-His3 were able to grow under selective conditions when the protein was stabilized.

Introduction

Selective protein degradation is essential for eukaryotic life, and altered protein degradation contributes to a number of medical conditions, including several types of cancer, neurodegenerative disease, cardiovascular disease, and cystic fibrosis^1-5. The ubiquitin-proteasome system (UPS), which catalyzes selective protein degradation, is an emerging therapeutic target for these conditions^6-10. Ubiquitin ligases covalently attach polymers of the 76-amino acid ubiquitin to proteins¹¹. Proteins that have been marked with polyubiquitin chains are recognized and proteolyzed by the ~2.5 megadalton 26S proteasome¹². Studies initiated in the model eukaryotic organism Saccharomyces cerevisiae (budding yeast) have been foundational in the elucidation of protein degradation mechanisms in eukaryotic cells. The first demonstrated physiological substrate of the UPS was the yeast transcriptional repressor MATα2^13,14, and many highly conserved components of the UPS were first identified or characterized in yeast (e.g. ^15-26). Discoveries made in this versatile and genetically tractable model organism are likely to continue to provide important insights into conserved mechanisms of ubiquitin-mediated degradation.

Recognition and degradation of most UPS substrates require the concerted action of multiple proteins. Therefore, an important goal in characterizing the regulated degradation of a given unstable protein is to determine the genetic requirements for proteolysis. Classical approaches (e.g. pulse-chase and cycloheximide-chase experiments²⁷) for monitoring protein degradation in mammalian or yeast cells are laborious and time-consuming. While these types of methodology provide highly sensitive means for detecting protein degradation, they are not suitable for rapid analysis of protein degradation or large-scale screening for mutations that prevent protein degradation. Here, a yeast growth-based assay for the rapid identification of genetic requirements for the degradation of unstable proteins is presented.

In the yeast growth-based method for analyzing protein degradation, an unstable protein of interest (or degradation signal) is fused, in frame, to a protein that is required for yeast growth under specific circumstances. The result is an artificial substrate that may serve as a powerful tool to determine the genetic requirements of protein degradation of the unstable protein of interest. Conveniently, most commonly used laboratory yeast strains harbor a panel of mutations in genes encoding metabolic enzymes involved in the biosynthesis of particular amino acids or nitrogenous bases (e.g. ^20,28-30). These enzymes are essential for cellular proliferation in the absence of exogenously provided metabolites in whose synthesis the enzymes participate. Such metabolic enzymes may thus function as growth-based reporters for the degradation of unstable proteins to which they are fused. The genetic requirements for protein degradation can be readily elucidated, since mutations that prevent proteolysis will allow cells harboring the degradation reporter to grow under selective conditions.

A growth advantage is an indirect indication that a particular mutation increases the abundance of the protein of interest. However, direct biochemical analysis is required to confirm that a mutation permits growth through increased protein levels rather than via indirect or artifactual causes. The effect of a mutation on protein abundance may be confirmed by western blot analysis of steady-state protein levels in cells that do and do not harbor the particular mutation. A method for the rapid and efficient extraction of yeast proteins (sequential incubation of yeast cells with sodium hydroxide and sample buffer) in a form suitable for analysis by western blotting is also presented³¹. Together, these experiments facilitate the rapid identification of candidate regulators of protein degradation.

Protocol

1. Yeast Growth Assay to Identify Candidate Mutants Defective in Protein Degradation

1. Transform wild-type and mutant yeast cells with a plasmid encoding an unstable protein fused, in frame, to a reporter metabolic enzyme.
2. Inoculate transformants in 5 ml of synthetic defined (SD) minimal medium that is selective for cells harboring plasmid molecules. Incubate overnight at 30 °C, rotating.
3. Measure the optical density at 600 nm (OD₆₀₀) of each overnight culture.
   NOTE: Following overnight incubation, cells in culture may be in either logarithmic or stationary growth phase but should have reached a minimal OD₆₀₀ of 0.2. Very slow-growing yeast strains may require incubation times longer than one night, or inoculation of a greater number of cells, as determined empirically.
4. Prepare six-fold serial dilutions of transformed yeast cells in a sterile 96-well plate, beginning with cells diluted to an OD₆₀₀ of 0.2. Place each yeast transformant to be assayed in a different row in the 96-well plate.
   1. For each transformant, calculate the volume of overnight culture required to dilute cells to an OD₆₀₀ of 0.2 in a final volume of 200 µl. Add this volume of overnight culture to the corresponding well in Column 1. Add the appropriate amount of sterile water to bring the volume to 200 µl.
   2. For each row of yeast, add 125 µl of sterile water to the wells in Columns 2, 3, and 4.
      NOTE: Individually wrapped sterile 96-well plates may be packaged with sterile lids. The lids may be used as reservoirs for the sterile water that is distributed in this step. This allows simultaneous transfer of sterile water to all wells in a given column with a multichannel pipettor.
   3. Mix the contents of the first column (yeast diluted to an OD₆₀₀ of 0.2) by pipetting up and down with a multichannel pipettor.
   4. Transfer 25 µl of yeast from Column 1 to Column 2, using a multichannel pipettor. Mix by pipetting up and down. Transfer 25 µl from Column 2 to Column 3, and 25 µl from Column 3 to Column 4 (mixing well at each step).
5. Mix each sample with a multichannel pipettor. Proceeding from most dilute to least dilute columns of yeast, pipette 4 µl of each sample onto two plates containing the appropriate selective medium. Use one plate with medium that maintains plasmid selection (this plate serves as a yeast spotting and growth control). Use a second plate with medium that selects for plasmid maintenance and expression of the unstable protein fused to the reporter enzyme. Because yeast settle rapidly, mix cells by pipetting up and down at regular intervals.
   NOTE: Drier plates will more readily absorb liquid than freshly prepared plates and are therefore recommended for these experiments. Damp plates may be dried by incubation at room temperature in low humidity for 1 – 2 days or shorter incubations in a laminar flow hood. Plates may dry unevenly if the laminar air flow is parallel to the bench. Use of a template makes it easier to spot yeast cells at regular distances. Two sample templates are provided in Figure 1. These may be printed, cut out, and affixed to the inside of a Petri dish lid.
6. Allow plates to dry on the bench top.
7. Incubate plates at 30 °C for 2 – 6 days.
8. Photograph each plate after incubation.

Figure 1. Templates for spotting yeast cells onto 100-mm agar plates. These templates may be used to facilitate spotting yeast at regular distances with a multichannel pipettor. Templates may be printed, cut out, and affixed to the inside of a Petri dish lid. Place Petri dish with growth medium inside lid with template affixed. Templates are marked with a notch to track orientation. It is recommended that plates used in growth assays be similarly marked with a notch to track orientation. Templates for spotting four (A) or five (B) serial dilutions of yeast cells are provided. Please click here to view a printable version of this figure with 100-mm templates.

2. Biochemical Confirmation of Yeast Growth Assay

1. Growth of Yeast Cells and Post-Alkaline Protein Extraction (modified from ³¹)
   1. Transform wild-type and mutant yeast cells with a plasmid encoding the unstable protein.
   2. Inoculate transformants in 5 ml of SD medium that is selective for cells harboring plasmid molecules. Incubate overnight at 30 °C, rotating.
   3. Measure the OD₆₀₀ of each overnight culture.
      NOTE: Following overnight incubation, cells may be in either logarithmic or stationary growth phase but should have reached an OD₆₀₀ that will permit dilution to an OD₆₀₀ of 0.2 in 10 ml fresh selective medium (step 2.1.4). Very slow-growing yeast strains may require incubation times longer than one night, or inoculation of a greater number of cells, as determined empirically.
   4. Dilute yeast cells to an OD₆₀₀ of 0.2 in 10 ml fresh selective medium.
   5. Continue to incubate cells at 30 °C, rotating or shaking, until cultures reach an OD₆₀₀ between 0.8 and 1.2 (i.e. are in mid-logarithmic growth).
      NOTE: If the unstable protein of interest is under the control of a regulatable promoter, the optimal timing of induction of protein expression and cell harvest may vary according to previous studies or empirical observations.
   6. Collect 2.5 OD₆₀₀ units of culture in a 15-ml conical tube by centrifugation at 5,000 x g for 5 min at room temperature. Remove supernatant by pipetting or aspiration.
      NOTE: One OD₆₀₀ unit is defined as the amount of yeast present in 1 ml of culture at OD₆₀₀ of 1.0. The volume of culture (in ml) required to harvest 2.5 OD₆₀₀ units (V) can be determined using the following equation: V = 2.5 OD₆₀₀ units / Measured OD₆₀₀
   7. Resuspend cells in 1 ml distilled water. Transfer suspended cells to a microcentrifuge tube.
   8. Pellet cells by centrifugation at 6,500 x g for 30 sec at room temperature. Remove supernatant by pipetting or aspiration.
   9. Resuspend cells in 100 µl distilled water by pipetting up and down or vortexing, and add 100 µl 0.2 M NaOH. Mix by pipetting up and down. Incubate samples for 5 min at room temperature.
   10. Pellet cells (most of which have not yet released proteins and are still viable) by centrifugation at 18,000 x g for 5 min. Remove supernatant by pipetting or aspiration.
   11. Resuspend pellet in 50 – 100 µl 1x Laemmli sample buffer, which will lyse cells, by pipetting up and down or vortexing.
       NOTE: Removal of the alkaline supernatant following centrifugation and subsequent resuspension of cells in Laemmli sample buffer extracts proteins at a pH compatible with sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) using a Tris-glycine running buffer system and western blotting.
   12. To fully denature proteins, incubate lysates at 95 °C for 5 min.
       NOTE: Aggregation-prone proteins (e.g. proteins with several transmembrane segments) may become insoluble when incubated at 95 °C. Therefore, lysates should be incubated at lower temperatures (e.g. 37 °C – 70 °C) for 10 – 30 min, as empirically determined, for the analysis of such proteins.
   13. Cool lysates by placing on ice for 5 min.
   14. Centrifuge lysates at 18,000 x g for 1 min at room temperature to pellet insoluble material. Separate the supernatant (solubilized extracted protein) by SDS-PAGE prior to subsequent western blot analysis (section 2.2). Alternatively, store lysates at -20 °C.
2. Representative Western Blotting Protocol
   1. Load empirically determined volume of lysates in an SDS-PAGE gel.
   2. Run gel at 200 V until dye front has reached the bottom of the gel.
   3. Transfer proteins from gel to polyvinylidene fluoride (PVDF) membrane by wet transfer at 20 V for 60 – 90 min at 4 °C.
   4. Block membrane by incubating in 5% skim milk in Tris-Buffered Saline (TBS), rocking, for 1 hr at room temperature or overnight at 4 °C.
   5. Decant blocking solution.
   6. Incubate membrane with primary antibody specific for protein of interest (or epitope tag thereof) in 1% skim milk in TBS with 0.1% Tween-20 (TBS/T) for 1 hr at room temperature, rocking.
   7. Decant antibody solution, and wash membrane 3 x 5 min with TBS/T at room temperature, rocking.
   8. Incubate membrane with appropriate fluorophore-conjugated secondary antibody in 1% skim milk in TBS/T for 1 hr at room temperature, rocking.
      NOTE: Because fluorophores are light-sensitive, dilutions of fluorophore-conjugated antibodies should be prepared in the dark. Additionally, incubation of membranes in the presence of fluorophore-conjugated antibodies should occur in lightproof containers. This can be accomplished by wrapping incubation trays in aluminum foil.
   9. Decant antibody solution, and wash membrane 3 x 5 min with TBS/T at room temperature, rocking.
   10. Acquire image of membrane using Li-Cor Odyssey CLx and Image Studio software (or comparable imaging equipment and software), according to manufacturer recommendations.
   11. After imaging membrane, incubate the membrane with a primary antibody specific for a loading control protein in 1% skim milk in TBS/T for 1 hr at room temperature, rocking.
   12. Decant antibody solution, and wash membrane 3 x 5 min with TBS/T at room temperature, rocking.
   13. Incubate membrane with appropriate fluorophore-conjugated secondary antibody in 1% skim milk in TBS/T for 1 hr at room temperature, rocking.
   14. Decant antibody solution, and wash membrane 3 x 5 min with TBS/T at room temperature, rocking.
   15. Acquire image of membrane using Li-Cor Odyssey CLx and Image Studio software (or comparable imaging equipment and software), according to manufacturer recommendations.

Representative Results

To illustrate this methodology, the His3 enzyme has been fused to the carboxy-terminus of the model endoplasmic reticulum (ER)-associated degradation (ERAD) substrate, Deg1-Sec62 (Figure 2A) to create Deg1-Sec62-His3 (Figure 3). Deg1-Sec62 represents a founding member of a novel class of ERAD substrates that are targeted following persistent, aberrant association with the translocon, the channel primarily responsible for moving proteins across the ER membrane^32-34. Such unstable proteins have provisionally been called ERAD-T (for translocon-associated) substrates. Previous studies indicate that, upon aberrant translocon engagement, Deg1-Sec62 is targeted for degradation by the Hrd1 ubiquitin ligase (Figure 2B-D)^32,34,35. Factors required for the degradation of other Hrd1 substrates appear to be dispensable for Deg1-Sec62 degradation, suggesting a novel degradation mechanism³². Under conditions of impaired lipid binding and prolonged translocon association, apolipoprotein B, the protein component of mammalian low-density lipoproteins, appears to be degraded by a related mechanism^36-38. Therefore, Deg1-Sec62 may provide a useful model for degradation of medically relevant translocon-associated proteins.

Wild-type and hrd1Δ yeast cells that lack the chromosomal HIS3 gene were transformed with an empty vector³⁹ or a plasmid encoding Deg1-Sec62-His3 and spotted onto selective growth medium (Figure 4). To confirm that equal numbers of transformed yeast cells were transferred to plates, cells were spotted onto medium lacking tryptophan (which selects for cells harboring the plasmid) but containing histidine. Similar growth was observed for all transformed yeast cells. Cells that express Deg1-Sec62-His3 were expected to grow in the absence of histidine only when the fusion protein is stabilized (i.e. when ERAD-T is compromised). Indeed, hrd1Δ yeast expressing Deg1-Sec62-His3 exhibited a growth advantage relative to wild-type cells expressing Deg1-Sec62-His3 on medium lacking tryptophan and histidine. However, marked fusion-protein-dependent growth in the absence of histidine was observed even in the presence of Hrd1. In order to increase the stringency of the assay, medium lacking histidine was supplemented with 3-amino-1H-1,2,3-triazole (3-AT), a competitive inhibitor of the His3 enzyme⁴⁰. Yeast expressing Hrd1 grew very poorly on medium lacking histidine supplemented with 1 – 2 mM 3-AT; when HRD1 was deleted, cell growth was restored. Inclusion of 3-AT at a concentration of 3 mM dramatically inhibited growth of all cells, regardless of the presence or absence of Hrd1. These results are consistent with Hrd1-dependent substrate degradation.

Next, the steady-state abundance of the Deg1-Sec62-His3 protein in yeast expressing or lacking the Hrd1 enzyme was directly tested. Western blotting analysis indicated a comparable increase in Deg1-Sec62-His3 and Deg1-Sec62 protein in hrd1Δ yeast relative to wild-type cells (Figure 5). This confirms a role for Hrd1 in the regulation of levels of both proteins. Hrd1-dependent degradation of Deg1-Sec62 proceeds after the protein aberrantly engages the ER translocon³². Importantly, Deg1-Sec62-His3 aberrantly engages the translocon in a similar manner (unpublished data), further validating the use of Deg1-Sec62-His3 as a growth-based reporter for degradation of Deg1-Sec62 specifically and translocon-associated proteins generally.

Figure 2: Model for degradation of Deg1-Sec62 following aberrant translocon engagement. (A) Schematic depiction of Deg1-Sec62. Deg1 (the amino-terminal 67 amino acids from MATα2) is followed, in sequence, by the Flag (F) epitope, the 2-transmembrane endoplasmic reticulum (ER) protein Sec62, and two copies of the S. aureus Protein A (PrA). For clarity, the fusion protein is referred to as Deg1-Sec62. (B) Following normal insertion of its two transmembrane segments into the ER membrane, persistent interaction of Deg1-Sec62 with the translocon triggers abnormal, Deg1-dependent translocon engagement. A portion of the initially cytosolic amino-terminal tail aberrantly enters—and likely remains within—the translocon. (C) Following abnormal translocon engagement, Hrd1 recognizes and ubiquitylates Deg1-Sec62. Red circles indicate ubiquitin molecules. (D) Deg1-Sec62 is then extracted from the ER membrane and degraded by the proteasome, likely relieving translocon obstruction.

Figure 3: Schematic depiction of Deg1-Sec62-His3 following aberrant translocon engagement. Deg1 is followed, in sequence, by the Flag (F) epitope, the 2-transmembrane ER protein Sec62, two copies of the S. aureus Protein A (PrA), and the yeast His3 enzyme. For clarity, the fusion protein is referred to as Deg1-Sec62-His3.

Figure 4: Fusing His3 to Deg1-Sec62 permits selection of degradation-defective mutants. Serial dilutions of wild-type (HRD1) and hrd1Δ yeast transformed with an empty vector or a plasmid encoding Deg1-Sec62-His3 were spotted onto medium lacking tryptophan, medium lacking tryptophan and histidine, and medium lacking tryptophan and histidine supplemented with 3-amino-1H-1,2,3-triazole (3-AT), a competitive inhibitor of His3, at the indicated concentrations. Please click here to view a larger version of this figure.

Figure 5: Increased abundance of Deg1-Sec62 and Deg1-Sec62-His3 in cells lacking Hrd1. Protein extracts were prepared from wild-type (+) and hrd1Δ(Δ) yeast expressing Deg1-Sec62 or Deg1-Sec62-His3. Proteins (equivalent to 0.125 OD₆₀₀ units) were separated by SDS-PAGE, followed by western blotting with rabbit anti-mouse secondary antibodies, which directly bind the Protein A epitopes of the fusion proteins. Subsequent western blotting with antibodies specific to Pgk1 provides a loading control.

Solution	Components	Comments
Synthetic Defined (SD) Minimal Yeast Medium	2 % dextrose, 0.67 % yeast nitrogen base without amino acids, 0.002 % adenine, 0.004 % uracil, 0.002 % arginine, 0.001 % histidine, 0.006 % isoleucine, 0.006 % leucine, 0.004 % lysine, 0.001 % methionine, 0.006 % phenylalanine, 0.005 % threonine, 0.004 % tryptophan. For solid (plate) medium, include 2 % agar.	1. Selective medium is prepared by omitting appropriate amino acid(s) or nitrogenous bases.
		2. For convenience, these ingredients may be maintained as concentrated stock solutions as follows. Amino acids may be maintained as 100X stock solution containing all desired amino acids. Yeast nitrogen base may be maintained in a 20X stock solution (13.4 %). Dextrose may be maintained in a 40 % stock solution. Adenine and uracil may be maintained as 1 % stock solutions in 0.1 M NaOH.
		3. Sterilize medium by autoclaving.
1X Laemmli Sample Buffer	2 % SDS, 10 % glycerol, 5 % β-mercaptoethanol, 60 mM Tris HCl pH 6.8, 0.008 % bromophenol blue	1. 1X Sample buffer is often prepared by diluting a more concentrated (e.g. 5X) stock.
		2. The dye bromophenol blue may be added to desired intensity. A "pinch" (very small amount tapped from the edge of a spatula) is typically sufficient.
0.2 M Sodium Hydroxide		Prepare in water. Sodium hydroxide reacts with glass. Therefore, for long-term storage, 0.2 M sodium hydroxide should be maintained in plastic containers.
Laemmli Running Buffer (5X)	125 mM Tris, 960 mM glycine, 0.5 % SDS	To prepare 1 L of 1X Laemmli Running Buffer, dilute 1:5 in dH2O
Tris Acetate-SDS Transfer Buffer (5X)	125 mM Tris acetate (pH 8.8), 960 mM glycine, 0.05 % SDS	To prepare 20 L of 1X Tris Acetate-SDS Transfer Buffer, combine 4 L of 5X stock, 4 L of methanol, and 12 L of dH2O
10X Tris-Buffered Saline (TBS)	500 mM Tris, 1.5 M NaCl; pH adjusted to 7.5	To prepare 1 L of 1X TBS, dilute 1:10 in dH2O. 1X TBS may be supplemented with the detergent Tween-20 and powdered skim milk, as appropriate.

Table 1: Solutions and buffers used in this study.

Strain Name	Alias	Relevant Genotype	Figures	Source
VJY6	MHY500	MATa	4 and 5	Chen et al., 1993
		his3-Δ200
		leu2-3,112
		ura3-52
		lys2-801
		trp1-1
		gal2
VJY10		MATa	4 and 5	This study
		his3-Δ200
		leu2-3,112
		ura3-52
		lys2-801
		trp1-1
		gal2
		hrd1::kanMX4

Table 2: Yeast strains used in this study. Details of construction are available upon request.

Plasmid Number	Full Plasmid Name	Figure	Source
pVJ30	pRS414-PMET25-Deg1-Flag-Sec62-2xProtA	5	Rubenstein et al., 2012
pVJ121	pRS414-PMET25 (empty vector with MET25 promoter)	4	Mumberg et al., 1994
pVJ467	pRS414-PMET25-Deg1-Flag-Sec62-2xProtA-His3	5	This study
pVJ477	pRS414-PGAL4-Deg1-Flag-Sec62-2xProtA-His3	4	This study

Table 3: Plasmids used in this study. Note that all plasmids contain a yeast centromere to allow replication in yeast cells, the TRP1 gene for selection in yeast cells, and the AmpR gene for maintenance in bacterial cells. Plasmid maps, sequences, and details of construction are available upon request.

Discussion

The methodology presented here allows for the rapid determination and biochemical confirmation of genetic requirements for protein degradation in yeast cells. These experiments highlight the utility and power of yeast as a model eukaryotic organism (several excellent reviews of yeast biology and compilations of protocols for handling, storing, and manipulating yeast cells (e.g. ^41-44) are available for investigators new to the organism). The techniques can readily be applied to investigate the degradation and abundance of a variety of classes of proteins. For example, others have employed this strategy to characterize the degradation mechanisms of unstable cytosolic, nuclear, and ER luminal and transmembrane proteins^45-49.

A few factors must be considered in the choice of metabolic enzyme to fuse to an unstable protein. First, it is essential that a functional version of the gene encoding the enzyme not be present in the host genome. To minimize false positive results (i.e. growth under selective conditions when the unstable protein is actually degraded), it is recommended to work with strains that harbor non-reverting mutant alleles of the reporter gene (preferably complete gene deletions)⁵⁰. Another consideration for reporter enzyme selection is the availability of competitive inhibitors, which can be included in the selective growth medium to reduce background growth and enhance assay stringency. This may be useful in cases of proteins with relatively low turnover rates even in the presence of fully functional degradation mechanisms. In the representative experiments presented here, the inclusion of 3-AT, which competitively inhibits the His3 enzyme, reduces background growth⁴⁰. Similarly, the compound 6-azauracil inhibits Ura3, an enzyme required for uracil biosynthesis⁵¹. The inhibitor concentration at which growth occurs in degradation-defective, but not wild-type, cells must be determined empirically. Some metabolic enzymes may also be counter-selected against. Under counter-selective conditions, cells may grow only when the unstable protein is degraded (and the fused metabolic enzyme is not present). For example, Ura3 converts the compound 5-fluoroorotic acid (5-FOA) to the toxic compound, 5-fluorouracil⁵². Cells expressing a Ura3-fusion protein will only grow in the presence of 5-FOA if the Ura3 fusion protein is degraded. Similarly, the compound 5-fluoroanthranilic acid (5-FAA) is toxic to cells with a functional tryptophan biosynthesis pathway. 5-FAA may thus be used to counter-select for cells expressing Trp1-fusion proteins⁵³. Counter-selection strategies may be useful for the identification of suppressors of degradation-impairing mutations.

The promoter used to drive expression of a degradation reporter must also be carefully selected. As low levels of biosynthetic enzymes may be sufficient to support growth in the absence of exogenously supplied metabolite, a weak promoter is recommended⁵⁴. In the representative experiments presented here, the GAL4 promoter, which is repressed in the presence of glucose⁵⁵, is used to promote the transcription of Deg1-Sec62-His3. Basal expression of this fusion protein under repressing conditions (i.e. 2% glucose) is sufficient to support growth under selective conditions (i.e. absence of histidine and presence of 1 – 2 mM 3-AT) when the degradation mechanism is disabled. However, protein levels sufficient to support growth under selective conditions are likely to be below the threshold of detection by western blotting. Therefore, it may be necessary to drive expression with a weak promoter for the growth assay and a stronger promoter for biochemical confirmation. In the case of Deg1-Sec62 (with or without the His3 fusion), a more robust promoter (here, the MET25 promoter³⁹) is required for protein visualization by western analysis.

As described here, the yeast-based growth assay may be performed on a small-scale, candidate-based approach. Serial dilutions of yeast cells are prepared in a 96-well plate and transferred by pipetting to solid growth medium; an alternative method for the efficient and reproducible transfer of diluted yeast cell suspensions onto solid medium is the use of a multi-pin replicator, commonly referred to as a “frogger”⁵⁶. The ideal time to photograph plates will vary with yeast strains and conditions. It is recommended to photograph a given plate when colonies from the fastest growing culture first become visible at the most dilute spot. This is typically the point at which differences in growth rates among samples are most obvious. It may be advisable to take photographs on multiple days, particularly in the case of yeast exhibiting a wide range of growth rates.

The growth-based reporter assay may also be adapted for large-scale analyses. For example, a degradation reporter may be introduced into a commercially available library of ~5,000 viable haploid yeast gene deletion strains using Synthetic Genetic Array (SGA) technology^46,57. In this technique, a haploid yeast strain with a chromosomally integrated metabolic reporter fusion protein is mated to each strain of the gene deletion library. The resultant diploid cells are induced to sporulate (undergo meiosis) and subjected to selection for haploid meiotic progeny harboring both the metabolic reporter and individual gene deletions. These strains are then transferred en masse to medium selective for cells in which the protein has been stabilized. As for small-scale analyses, if a gene with a role in protein degradation is deleted, the fusion protein will be stabilized, and cell growth will be enhanced. Comparable approaches have been devised that allow for simultaneous transformation of a large collection of strains with extra-chromosomally maintained plasmids; this strategy obviates chromosomal integration, mating, sporulation, and meiotic selection⁵⁴.

When a mutation is found to confer a growth advantage to cells harboring a metabolic reporter, it is necessary to biochemically confirm that the mutation increases the abundance of the protein of interest. A rapid and reliable yeast lysis procedure, closely adapted from the method of Kuhsnirov³¹, is presented. This protocol allows for the extraction of proteins in a form directly suitable for analysis by western blotting. For the analysis of a given protein, the amount of lysate to be loaded per well, acrylamide gel properties, choice of membrane, antibodies used and dilutions thereof, and method of detection must be empirically determined. The representative western blotting protocol described utilizes secondary antibodies conjugated to fluorescent dyes; other commonly used protocols rely on chemiluminescence dependent on antibody-conjugated enzymes⁵⁸. As described here, the membrane used to detect the protein of interest may be directly reprobed with an antibody for a loading control protein. If the primary antibodies used to detect the protein of interest and loading control protein have been raised from the same species, reprobing the membrane is possible as long as the bands arising from these proteins do not co-migrate. If, however, the primary antibodies used to detect the protein of interest and loading control protein have been raised from different species, the same membrane may be sequentially probed, even if bands co-migrate, if the secondary antibodies have been conjugated to fluorophores with different emission wavelengths. In the case that the protein of interest and loading control protein co-migrate and the respective primary antibodies have been raised from the same species, samples may be resolved on two SDS-PAGE gels, transferred to PVDF membrane, and probed separately with antibodies specific for the protein of interest and loading control protein. Alternatively, loading consistency may be judged by incubating membranes with non-specific protein stains (e.g. Coomassie or Ponceau S). Further, the representative western blotting protocol assumes a protein or epitope that is detected by sequential incubation of primary and secondary antibodies, as is typical. The fusion proteins analyzed in the representative results contain two epitopes derived from Staphylococcus aureus Protein A (PrA in Figures 2 and 3). Protein A binds directly to mammalian immunoglobulins and therefore can be detected using secondary antibody alone (i.e. no primary antibody incubation step is required)⁵⁹. It is possible that fusion of a reporter enzyme may influence protein abundance or degradation. It is therefore advisable to biochemically confirm the results using a version of the substrate unencumbered by the reporter protein. Finally, both assays described here rely on differences in steady-state protein levels as a proxy for differences in protein stability. Because protein abundance reflects the integration of rates of protein synthesis and degradation, further biochemical analysis (e.g. cycloheximide chase or pulse chase experiments) must be employed to directly analyze a protein’s dynamic degradation profile.

The representative results establish a novel application of this protocol for the determination of genetic requirements for degradation of a protein that aberrantly engages the ER translocon. Deg1-Sec62-His3 conferred a yeast growth advantage under selective conditions when the degradation pathway was inactivated (i.e. in the absence of the Hrd1 ubiquitin ligase). A fast and reliable protein extraction method followed by western blotting confirmed an increase in abundance of Deg1-Sec62 (with or without His3) in the absence of Hrd1. Previous studies indicate that the mechanism of Hrd1-dependent degradation of translocon-associated proteins differs from those of other Hrd1 ER luminal or transmembrane substrates³². Future work will employ the Deg1-Sec62-His3 fusion protein in large-scale genetic screens to identify novel genes required for this unique degradation mechanism.

Materials

Name	Company	Catalog Number	Comments
Desired yeast strains, plasmids, standard medium and buffer components			Yeast strains with desired mutations may be generated in the investigator's laboratory. Wild-type yeast and a variety of mutants are also commercially available (e.g. from GE Healthcare). Plasmids encoding fusion proteins may be generated in the investigator's laboratory.
3-amino-1H-1,2,4-triazole	Fisher Scientific	AC264571000	Competitive inhibitor of His3 enzyme. May be included in medium to increase stringency of growth assay using His3 reporter constructs.
Endoglycosidase H (recombinant form from Streptomyces plicatus)	Roche	11088726001	May be used to assess N-glycosylation of proteins; compatible with SDS and beta-mercaptoethanol concentrations found in 1x Laemmli sample buffer.
Disposable borosilicate glass tubes	Fisher Scientific	14-961-32	Available from a variety of manufacturers
Temperature-regulated incubator (e.g. Heratherm Incubator Model IMH180)	Dot Scientific	51028068	Available from a variety of manufacturers
New Brunswick Interchangeable Drum for 18 mm tubes (tube roller)	New Brunswick	M1053-0450	Tube roller is recommended to maintain overnight yeast starter cultures of yeast cells in suspension. A platform shaker or tube roller may be used to maintain larger cultures in suspension.
New Brunswick TC-7 Roller Drum 120V 50/60 H	New Brunswick	M1053-4004	For use with tube roller
SmartSpec Plus Spectrophotometer	Bio-Rad	170-2525	Available from a variety of manufacturers
Sterile 96-well flat bottom microtest plates with lid individually wrapped	Sarstedt	82.1581.001	Available from a variety of manufacturers
Pipetman Neo P8x20N, 2-20 μl	Gilson	F14401	Available from a variety of manufacturers

Name	Company	Catalog Number	Comments
Pipetman Neo P8x200N, 20-200 μl	Gilson	F14403	Single-channel and multichannel pipettors are used at various stages of the protocol. While multichannel pipettors reduce the pipetting burden at several steps, single-channel pipettors may be used throughout the entire protocol. Available from a variety of manufacturers.
Centrifuge 5430	Eppendorf	5427 000.216	Rotor that is sold with unit holds 1.5 and 2.0 ml microcentrifuge tubes. Rotor may be swapped for one that holds 15 ml and 50 ml conical tubes.
Plate imaging system (e.g. Gel Doc XR+ System)	Bio-Rad	170-8195	A variety of systems may be used to image plates, including sophisticated imaging systems, computer scanners, and camera phones.
Fixed-Angle Rotor F-35-6-30 with Lid and Adapters for Centrifuge Model 5430/R, 15/50 ml Conical Tubes, 6-Place	Eppendorf	F-35-6-30
15 ml screen printed screw cap tube 17 x 20 mm conical, polypropylene	Sarstedt	62.554.205	Available from a variety of manufacturers
1.5 ml flex-tube, PCR clean, Natural microcentrifuge tubes	Eppendorf	22364120	Available from a variety of manufacturers
Analog Dri-Bath Heater	Fisher Scientific	1172011AQ	Boiling water bath with hot plate may also be used to denature proteins
SDS-PAGE running and transfer apparatuses, power supplies, and imaging equipment or darkrooms for SDS-PAGE and transfer to membrane			Will vary by lab and application
Western blot imaging system (e.g. Li-Cor Odyssey CLx scanner and Image Studio Software)	Li-Cor	9140-01	Will vary by lab and application
EMD Millipore Immobilon PVDF Transfer Membranes	Fisher Scientific	IPFL00010	Will vary by lab and application
Primary antibodies (e.g. Phosphoglycerate Kinase (Pgk1) Monoclonal antibody, mouse (clone 22C5D8))	Life Technologies	459250	Will vary by lab and application
Secondary antibodies (e.g. Alexa-Fluor 680 Rabbit Anti-Mouse IgG (H+L))	Life Technologies	A-21065	Will vary by lab and application