Determining Atg protein stoichiometry at the phagophore assembly site by fluorescence microscopy

Abstract

In eukaryotic cells, autophagy is a lysosomal/vacuolar degradative pathway necessary for the turnover of different macromolecules. Autophagy is under precise regulation, not only qualitatively but also quantitatively, and excess or reduced levels of autophagy may lead to various human diseases. In yeast, genetic screens led to the identification of more than 30 autophagy-related (ATG) genes, and most of the gene products reside at the phagophore assembly site (PAS). However, our attempt to understand the quantitative properties of autophagy is usually hampered, because traditional methods of analysis cannot provide stoichiometric information. We have recently used a fluorescence microscopy-based method to study the stoichiometry of Atg proteins at the PAS, trying to explain the mechanism of how the vesicle formation process is precisely regulated. This article describes a practical guide on this method. Its application and further analysis will improve our understanding of the quantitative properties of autophagy.

1. Introduction

Ever since its initial discovery, the green fluorescent protein (GFP) has substantially changed the research methodology of modern biology. By fusing GFP and its various derivatives to other interesting, but otherwise invisible, proteins, researchers can “watch” the positions, movements and interactions of tagged proteins in different conditions. The application of fluorescent protein methodologies is still expanding. Recently, Wu and Pollard reported that protein concentration can be directly measured by fluorescence microscopy in Schizosaccharomyces pombe.¹ Inspired by their results, we applied similar methods in Saccharomyces cerevisiae to study the stoichiometry of Atg proteins during the vesicle formation process.²

This method is feasible and useful in the study of autophagy. First, most Atg proteins localize to a very restricted region, the phagophore assembly site (PAS, also known as the pre-autophagosomal structure).^3,4 This property makes the quantification process much easier. Second, it is also important to know the stoichiometry of Atg proteins at this locus. The PAS is thought to be the organization center for the formation of the Cvt vesicle and autophagosome. Multiple lines of evidence have pointed out that the amount of some Atg proteins (e.g., Atg8 and Atg9) at the PAS is under precise regulation and related to the quantitative property of the vesicle formation process, such as vesicle size or vesicle number.^5-7 However, traditional methods cannot provide information on the quantitative analysis of Atg proteins at a specific subcellular locus. In contrast, this fluorescence microscopy method can directly measure the protein amount in living cells and even in a time-course experiment.

In this protocol article, we describe the method to quantify the amount of Atg proteins at the PAS. Theoretically we can use this method to measure the protein amount at any subcellular localization. In our analysis of Atg proteins, the PAS is the clearest structure for fluorescence quantification. The method can be divided into two basic parts. The first part is to establish a standard curve showing the linear relationship between fluorescence intensity and protein amount; the second part is to use this standard curve to measure and calculate the protein amount at the PAS. In some cases, relative quantification provides sufficient information, for example, in asking how does the Atg8 concentration at the PAS change in different conditions, or what are the kinetics of Atg8 recruitment to the PAS? In this situation, after confirming the linear relationship between the protein amount and fluorescence intensity, it is possible to simply measure the relative intensity and convert it into relative protein amount without determining the absolute value.

2. Materials

2.1. Cells and culture

1. Cells: Choose a set of proteins with different expression levels and tag them to GFP, e.g., A-GFP, B-GFP, C-GFP, etc. (see Notes 1–3), which will be used to set up the standard curve. For the proteins of interest, also fuse them to GFP.

2. Growth medium (YPD): 1% yeast extract, 2% peptone and 2% glucose. This medium is stable for months at room temperature.

3. Synthetic minimal medium (SMD): 0.67% yeast nitrogen base, 2% glucose and auxotrophic amino acids and vitamins as needed. This medium is stable for months at room temperature.

4. Starvation medium (SD-N): 0.17% yeast nitrogen base without ammonium sulfate or amino acids, containing 2% glucose. This medium is stable for months at room temperature.

2.2. Standard protein: GST-GFP

1. Clone the ORF of GFP (S65T) into pGEX-4T-1 (GE Healthcare, 27-4580-01) and express it in E. coli strain BL-21. Follow a standard protein purification protocol to purify the fusion protein on glutathione-Sepharose beads (GE Healthcare, 17-0756-01).

2. Measure the concentration of purified protein (in ng/μl) using the bicinchoninic acid (BCA) assay (Pierce Chemical Co., 23223 and 23224) and convert the concentration to mol/μl ([mol/μl] = [ng/μl]*10^-9/MW_GST-GFP, MW_GST-GFP= 54,808.7).

2.3. Microscopy

1. Samples are examined with an inverted fluorescence microscope (DeltaVision Spectris; Applied Precision, LLC or equivalent equipment).

2. Microscopy images are analyzed and quantified using softWoRx (Applied Precision, LLC) or equivalent software.

3. Concavity slides (Fisher, S175201) for time-course experiments.

2.4. Antibody

Living colors A.v. monoclonal antibody JL-8 (Clontech, 632381).

3. Methods

3.1. Standard curve: linear relationship between protein amount and fluorescence intensity

3.1.1. Measure fluorescence intensity per cell

1. Yeast strains expressing GFP-tagged proteins (A-GFP, B-GFP, C-GFP, etc.), are cultured in 5 ml YPD medium at 30°C in test tubes. Cells are grown to log phase, then diluted, and cells are then regrown to early log-phase (A₆₀₀ = 0.6–0.8). For experiments under starvation condition, cells are shifted to SD-N medium and cultured at 30°C for another 2 h.

2. Harvest 1 ml culture. Centrifuge at 3,000 rpm for 2 min to pellet the cells. If time permits, an additional aliquot of cells is collected to determine the amount of GFP fusion-protein per cell (see section 3.1.2).

3. The supernatant fraction is discarded. The cell pellet is washed with 1 ml SMD medium without vitamins (see Notes 4 and 5) and centrifuged again.

4. Discard the supernatant fraction and resuspend the pellet fraction in 30 μl SMD medium (see Note 5).

5. Mount 5 μl of sample on a microscope slide and cover it with a coverslip. Press the coverslip down gently.

6. Observe by fluorescence microscopy and take pictures. For each picture, take 12 Z-section images at a 0.5-μm interval (see Note 6).

7. Deconvolve the images.

8. In softWoRx software, go to “View → Quick Projection.” Input 12 Z-section images and stack them into a 2-D image using “sum projection.”

9. Open the projected image. Go to “Tool → Data Inspector.” Draw a circle with the proper size to cover the whole yeast cell. Record the “Total” intensity within this region.

10. Quantify the fluorescence intensity of wild-type cells in which no GFP-fusion protein is expressed. Use this value as the autofluorescence background. Subtract the background from the “Total” value obtained in the previous step.

11. For each strain (A-GFP, B-GFP, C-GFP, etc.) quantify 200 cells. Calculate the mean value and S.E.M. (standard error of the mean: standard deviation divided by the square root of the sample size).

3.1.2. Measure amount of GFP-fusion protein per cell

1. Culture cells (A-GFP, B-GFP, C-GFP, etc.) under the same condition as used for the microscopy experiments. Harvest 1 OD₆₀₀ unit of cells (equivalent to 1 ml of culture at A₆₀₀ = 1.0). Ideally, these samples should be collected at the same time that the cells are prepared for the microscopy analysis, using the same culture (section 3.1.1).

2. Resuspend in 10% TCA. Keep on ice for 30 min (the samples can also be stored at 4°C or -20°C for later processing).

3. Centrifuge at 13,000 rpm for 5 min at 4°C. Discard the supernatant fraction.

4. Add 1 ml ice-cold acetone. Resuspend by sonication, using a water bath sonicator to disrupt the pellet fraction.

5. Centrifuge at 13,000 rpm for 5 min at 4°C. Discard the supernatant fraction. Dry the pellet fraction on ice.

6. Add 50 μl 1× SDS sample buffer (equivalent to 0.02 OD₆₀₀ cell/μl) and 0.4–0.6 mm acid washed soda lime glass beads. Glass beads are prepared as described previously.⁸

7. Mix on a vortex at 4°C for 5 min. Incubate at 85°C for 5 min (see Note 7). Centrifuge at 13,000 rpm for 1 min at 4°C.

8. Load an appropriate amount of the cell lysate sample for SDS-PAGE. On the same gel, also load a concentration gradient of standard protein, GST-GFP (see Note 8).

9. Follow a standard protocol to do western blotting. Use YFP antibody and overnight incubation. After development with appropriate exposure (see Note 9), scan the film.

10. Quantify the band intensity using NIHimage (see Note 10). Compare the band intensity of the GFP fusion protein with the GST-GFP standard protein to determine the protein concentration in mol/OD₆₀₀ units of cells.

11. Repeat the western blot and quantification three times. Calculate the mean value and S.E.M.

12. Convert the protein concentration into molecules/cell by multiplying by Avogadro's number and dividing by the number of cells per OD₆₀₀.

3.1.3. Standard curve: fluorescence intensity vs. protein amount

1. For A-GFP, B-GFP, C-GFP, etc., plot the fluorescence intensity against the protein amount (X axis: molecules/cell; Y axis: intensity/cell).

2. Set intercept to 0 (see Note 11) and do linear regression. The slope corresponds to the ratio between fluorescence intensity and number of molecules.

3. Calculate R² to evaluate the fit of the line (see Note 12).

3.2. Use standard curve to calculate the concentration of Atg proteins at the PAS

1. Culture cells expressing the GFP fusion protein of interest. Choose the culture condition based on your specific aim.

2. Prepare microscopy sample and take pictures as described previously.

3. (Optional) In time-course experiments, prepare slides as follows: Add warm medium containing 1.8% agar into the concave region of a concavity slide. Cover it with a coverslip and wait until complete solidification of the agar. Remove the coverslip by “sliding” to get a smooth surface on the solidified agar. Mount your sample to this region and cover it with a new coverslip. Seal the edge of the coverslip with nail polish to avoid liquid evaporation. Take pictures at every time-point as described previously but with relatively short exposure times to minimize photobleaching. The effect of photobleaching can be determined empirically.²

4. Deconvolve the images and make 2-D projections.

5. Draw a circle around the PAS dot. Record the intensity value.

6. Move the circle to an adjacent cytosolic region. Record the intensity as background.

7. Subtract the background value from the raw PAS intensity.

8. Use the slope obtained from the standard curve to convert the fluorescence intensity into number of molecules (see Note 13).

9. Repeat this quantification process. Calculate the mean and S.E.M.

4. Notes

1. It is not necessary that A, B, C, etc., are different proteins as long as their expression level is different. For instance, in our initial study we used strains expressing GFP-Atg8 driven by various promoters.

2. It is critical that the range of your standard curve is wide enough to cover the abundance of the protein you want to study. Large scale data on gene expression level⁹ is available in the Saccharomyces Genome Database (www.yeastgenome.org), which can help you decide which protein to choose.

3. Chromosomal tagging is strongly suggested rather than using CEN plasmids because of the consistent expression level from cell to cell. If you tag the fluorophore to the C terminus of your protein, the PCR-based method can be used as described by Longtine et al.¹⁰ If C-terminal tagging is not feasible such as with Atg8, you should clone the N-terminal fusion protein into an integration plasmid and then integrate it into the yeast genome. In this case, the endogenous copy of the gene should be deleted to avoid changing the normal protein level. We found essentially identical results using either N- or C-terminal fusions; however, the function of the fusion protein should always be confirmed.

4. YPD medium produces a high autofluorescence background. For the same reason, do not add vitamins in SMD medium at this step.

5. Starved cells are washed and resuspended in SD-N medium to maintain the nutrient-deficient condition.

6. No more than three pictures should be taken on the same slide to minimize the effect of photobleaching. The total depth of each stack is 5.5 μm, which is the approximate diameter of a normal yeast cell. Focus to the middle plane of the cell.

7. For proteins such as Atg9, high temperature impairs its solubility. In such cases, incubate the protein samples at 55°C for 15 min.

8. It is critical to do pre-experiments in order to decide on the appropriate loading amount. To make sure the band intensity is in the linear range, use at least two concentrations of samples.

9. Prevent overexposure during development. Saturated bands significantly affect the quantification results.

10. Draw a rectangle big enough to cover one single western band. First, move the rectangle to a clear area on the film and measure the intensity, which is used for the background. Then move the rectangle to the band you want to quantify, measure the intensity and subtract the background. Other equivalent software such as ImageJ can also be used.

11. When the protein concentration is zero, we expect the fluorescence intensity to theoretically be zero as well.

12. The R² value varies from 0 to 1. An R² of 1.0 indicates that the regression line perfectly fits the data. There is no objective criteria to judge what value of R² is acceptable, although the R² values in similar studies showing the linearity between fluorescence intensity and protein amount are higher than 0.95.^1,2

13. One obstacle for fluorescence quantification is that the measured intensity usually varies in repetitive experiments. Therefore, every time you want to quantify the absolute amount of a protein, it is important to include two proteins used in determining the standard curve (e.g., the two with the highest and lowest protein level) in the assay to calibrate the slope of the standard curve.