Systematic analysis of assembly intermediates in yeast to decipher the mitoribosome assembly pathway

Abstract

Studies of yeast mitoribosome assembly have been historically hampered by the difficulty of generating mitoribosome protein-coding gene deletion strains with a stable mitochondrial genome. The identification of mitochondrial DNA stabilizing approaches allows for the generation of a complete set of yeast deletion strains covering all mitoribosome proteins and known assembly factors. These strains can be used to analyze the integrity and assembly state of mitoribosomes by determining the sedimentation profile of these structures by sucrose gradient centrifugation of mitochondrial extracts, coupled to mass spectrometry analysis of mitoribosome composition. Subsequent hierarchical cluster analysis of mitoribosome subassemblies accumulated in mutant strains reveals details regarding the order of protein association during the mitoribosome biogenetic process. These strains also allow the expression of truncated protein variants to probe the role of mitochondrion-specific protein extensions, the relevance of protein co-factors, or the importance of RNA-protein interactions in functional sites of the mitoribosome. In this chapter, we will detail the methodology involved in these studies.

1. Introduction

Ribosome biogenesis involves the processing and modification of ribosomal RNAs (rRNAs) coordinated with the incorporation of ribosomal proteins. For the ribosomes present in mitochondria (mitoribosomes), their assembly is complicated by the dual genetic origin, nuclear and mitochondrial, of their rRNA and protein (MRP) components [1]. Furthermore, mitoribosome composition and structure vary significantly among organisms [2], leading to species-specific assembly constraints. The biogenetic process requires numerous nucleus-encoded transacting factors, some of which are conserved in bacterial systems, in mitochondria across species, or species-specific, acting in all steps of the process.

Several approaches have been used to address aspects of the mitoribosome biogenesis pathway in yeast and other organisms. Searching for mitochondrial RNA-binding proteins in mammalian cells, or proteomics analyses of the mitoribosome interactome have allowed for the identification of potential mitoribosome assembly factors [3–7]. Also, studies in patients with mitochondrial translation efficiency disorders have allowed identifying new assembly factors and the relevance of MRPs for mitoribosome assembly across tissues [1, 8–10]. Other studies have leveraged the better characterized bacterial ribosome assembly pathway [11] and screens in yeast [12, 13] to identify potential mitoribosome assembly factors that were subsequently characterized in human cultured cells or mouse models [14–16]. Furthermore, structures of the mitochondrial ribosome from human HEK293 cells [17–23] and the human parasite Trypanosoma brucei [24–27] in native states of assembly have revealed insights into the timing of rRNA folding and protein incorporation during the final steps of ribosomal maturation.

Two approaches have so far been used to disclose the overall assembly pathway of the mitochondrial ribosome. First, a study performed in human HeLa cells analyzed the order of MRP assembly using stable pulse-chase isotope labeling in cell culture (SILAC) and mass spectrometry analysis of the human 55S mitochondrial monosome [28]. The approach is based on the model that the kinetics of incorporation of different MRPs into 55S mitoribosomes indicates their relative assembly order [28]. This study provided a useful albeit low-resolution draft of the assembly pathway by defining sets of early, intermediate, and late assembly proteins [28]. Second, a study performed by our group on the yeast Saccharomyces cerevisiae characterized a collection of strains systematically deleted for one of the mitochondrial large 54S subunit (mtLSU) proteins or known assembly factors [29]. The S. cerevisiae mitoribosome is a 74S ribonucleoprotein particle whose mtSSU is formed by a 15S rRNA and 34 MRPs [30], and its mtLSU by a 21S rRNA and 46 MRPs [31]. The two rRNAs and one of the mtSSU proteins, Var1 -renamed as uS3m- are encoded in the mitochondrial genome (mtDNA), whereas all other proteins are encoded in the nuclear genome. In our study, the mtLSU subassemblies that accumulated in each deletion strain were obtained by sucrose gradient sedimentation fractionation, their composition deciphered by mass-spectrometry, and their potential order in the assembly pathway established by clustering analysis. We are now applying this approach to the characterization of the yeast 37S small subunit (mtSSU) and will describe it in full detail in this chapter.

Studies of yeast mitoribosome assembly have been historically difficult due to the loss of mtDNA inherent to all strains defective in mitochondrial translation. The approach presented here takes advantage of the identification of multicopy suppressors of the mtDNA instability, which affords the possibility of generating mitoribosome deletion strains carrying mtDNA [32]. All the S. cerevisiae strains used were isogenic to the respiratory-robust wild-type strain W303I⁰ carrying intronless mtDNA to distinguish mitoribosome assembly defects due to altered processing of the intron-containing 21S rRNA gene. To prevent the mtDNA loss that occurs in yeast mitoribosome assembly mutants [29], all strains are engineered to overexpress a previously identified suppressor of this phenotype (the ribonucleotide reductase catalytic subunit RNR1) [32, 33]. Furthermore, since the mtDNA-encoded mtSSU protein Var1 will not be synthesized when mitochondrial translation is impaired, which can be a confounding variable, all the strains were transformed with a construct that successfully relocates a recoded version of the VAR1 gene (VAR1^U) to the nucleus as described [34].

The systematic analysis of mitoribosome assembly intermediates in deletion strains described in this chapter is limited by the potential heterogeneity of the subassemblies analyzed in each strain. This approach does not distinguish among “on pathway” assembled intermediates, “dead-end” aberrant subassemblies, and potential artifact subassemblies resulting from fragmentation of larger assemblies during experimental manipulation. The global survey of mitoribosome mutant yeast strains involved in this approach provides a strong model for mitoribosome subunit assembly that will serve as a framework for further pathway refinement studies in wild-type cells.

2. Materials

Prepare all solutions using analytical grade reagents and ultrapure water obtained by deionizing distilled water to a resistivity of 18 MΩ-cm at 25 ^oC. Store solutions at room temperature (RT, 25 °C) unless otherwise indicated.

2.1. Generation of a Collection of Yeast Strains KO for Genes Coding for Mitoribosome Proteins and Assembly Factors

2.1.1. Amplification of Gene-Targeted KanMX4 Deletion Cassettes

1. Agarose.

2. 1X TBE: 100 mM Tris-HCl, 100 mM boric acid, 2 mM ethylenediaminetetraacetic acid (EDTA) pH 8.0.

3. DNA Extraction kit.

2.1.2. Yeast Transformation

1. Yeast strain a/α W303I⁰ + VAR1^U + RNR1 (genotype: diploid, ade2–1 his3–11,15 leu2–3,112 trp1–1 ura3–1, URA3::pRS316-VAR1^U, LEU2::YEplac181-RNR1, ρ⁺ I⁰) [29].

2. Synthetic growth media: 0.67 % (w/v) yeast nitrogenous base without amino acids (WO), 2 % (w/v) glucose. Add 2% (w/v) agar for solid media. Sterilize by autoclaving.

3. AHW – 50x auxotrophic markers: 2 mg/mL adenine, 2.5 mg/mL histidine, 2.5 mg/mL tryptophan. Dissolve 1g of adenine, 1.25 g histidine, and 1.25 g tryptophan in 400 mL of water and bring up to a final volume of 500 mL (see Note 1). Sterilize by filtration through a 0.2 μm filter and store at RT in either a sterile light-sensitive amber bottle or wrap the bottle in aluminum foil as tryptophan is photosensitive.

4. TEL solution: 10 mM Tris-HCl pH 7.5, 1 mM EDTA, 100 mM lithium acetate. Sterilize by filtration through a 0.2 μm filter and store at RT.

5. 10 mg/mL sheared and denatured salmon sperm carrier DNA.

6. 40 % PEG-TEL solution: 40 % (w/v) polyethylene glycol (PEG 3500 or 4000), 10 mM Tris-HCl pH 7.5, 1 mM EDTA, 100 mM lithium acetate. Sterilize by filtration through a 0.45 μm filter and store at RT.

7. YPD liquid growth media: 1 % (w/v) yeast extract, 2 % (w/v) peptone, 2 % (w/v) glucose. Sterilize by autoclaving.

8. 50 mg/ml Geneticin (G418). Dissolve in water, sterilize by filtration through a 0.2 μm syringe tip filter and store at 4 °C.

9. 2.1.3 Genomic DNA Extraction and Validation of KanMX4 Cassette Integration

10. Haploid yeast KO strain in the W303I⁰ + VAR1^U + RNR1 background. For example, strain αW303-I⁰ ΔuL1 + VAR1^U + RNR1 (genotype MATα, ade2–1 his3–11,15 leu2–3,112 trp1–1 ura3–1, URA3::pRS316-VAR1, LEU2::YEplac181-RNR1, ΔuL1::KanMX, ρ⁺ I⁰)[29].

11. Solution A: 50 mM Tris-HCl pH 7.5, 10 mM EDTA pH 8.0, 0.3 % (v/v) β-mercaptoethanol, 0.5 mg/mL 100T Zymolyase.

12. 10 % (w/v) sodium dodecyl sulfate (SDS).

13. 8 M ammonium acetate.

14. Isopropanol.

15. 80 % (v/v) ethanol.

2.2. Sucrose Gradient Analysis to Establish the Mitoribosome Profile and Detect Assembly Intermediates

2.2.1. Growth of Yeast Cells

1. WO-Gal (low-glucose) synthetic growth media: 0.67 % (w/v) yeast nitrogenous base without amino acids, 2 % (w/v) galactose, 0.5 % (w/v) glucose. Dissolve 6.71 g of yeast nitrogenous base without amino acids, 20 g of galactose, and 5 g of glucose in 900 mL of water. Adjust pH to 5.8–6.0 with potassium hydroxide and bring up to a final volume of 1 L with water. For solid media, additionally, dissolve 20 g/L (2 % w/v) of agar. Sterilize by autoclaving (see Note 2). For WO-Gal-AHW, add 20 mL of 50x AHW per 1 L of sterile WO-Gal to achieve 1x working strength.

2.2.2. Isolation of Mitochondria

1. Tris-DTT buffer: 100 mM Tris-HCl pH 8.8, 10 mM dithiothreitol (DTT).

2. 1.2 M Sorbitol: Dissolve 874.416 g of sorbitol in 3 L of water. Bring to a final volume of 4 L. To remove traces of metals and other ionic contaminants, deionize sorbitol overnight at RT with AG 501-X8 resin from Bio-Rad (Hercules, CA). Filter through a 0.2 μm filter and store at 4 °C.

3. Digestion buffer: 1.2 M sorbitol, 20 mM potassium phosphate pH 7.4 (see Note 3), 0.25 mg/mL 100T zymolyase.

4. Homogenization buffer: 10 mM Tris-HCl pH 7.5, 1 mM EDTA pH 8.0, 0.2 % (w/v) bovine serum albumin (BSA), 1 mM phenylmethylsulfonyl fluoride (PMSF), 0.6 M sorbitol. Prepare fresh before use.

5. SH buffer: 0.6 M sorbitol, 20 mM HEPES pH 7.4. Sterilize by filtration through a 0.2 μm filter and store at 4 °C.

6. Floor centrifuge capable of reaching 14,000 xg at 4 °C.

7. 50 mL glass dounce with a loose Teflon pestle.

8. 10 mL glass dounce with a tight Teflon pestle.

2.2.3. Preparation of a 10–30 % Linear Sucrose Gradient

1. 10 % sucrose gradient buffer: 10 mM Tris pH 7.4, 100 mM NH₄Cl, 10 mM MgCl₂, 1 mM PMSF, 0.05 % (w/v) digitonin (see Note 4), 1x yeast protease inhibitor cocktail, 0.1 U/μL RNase inhibitor, 10 % (w/v) sucrose (see Note 5).

2. 30 % sucrose gradient buffer: 10 mM Tris pH 7.4, 100 mM NH₄Cl, 10 mM MgCl₂, 1 mM PMSF, 0.05 % (w/v) digitonin, 1x yeast protease inhibitor cocktail, 0.1 U/μL RNase inhibitor, 30 % (w/v) sucrose.

3. Polypropylene ultracentrifuge tubes (13 × 51 mm).

4. Biocomp Gradient Master with marker block, layering cannula, MagnaBase tube holder, 4 mm short tube caps and leveling tool.

2.2.4. Mitoribosome Extraction and Sedimentation Analysis

1. Extraction buffer: 10 mM Tris-HCl pH 7.4, 100 mM NH₄Cl, 10 mM MgCl₂, 1 mM PMSF, 0.25 mM DTT, 0.6 % (w/v) high purity digitonin (see Note 4), 1x yeast protease inhibitor cocktail, 0.1 U/μL RNase inhibitor.

2. Ultracentrifuge capable of reaching 200,000 xg.

3. Swinging-bucket rotor for ultracentrifuge (i.e., SW55ti).

4. 4x LB (4x Laemmli sample buffer): 200 mM Tris-HCl pH 6.8, 4 % (w/v) SDS, 40 % (v/v) glycerol, 4 % (v/v) β-mercaptoethanol, 0.05 % (w/v) bromophenol blue.

2.2.5. SDS-PAGE and Immunoblot Analysis of Sedimented Mitoribosomal Particles

1. 12 % SDS-PAGE polyacrylamide gel: Gels are made in house from a 12 % resolving gel (12 % / 0.32 % (w/v) acrylamide/bis-acrylamide, 375 mM Tris-HCl pH 8.8, 0.1 % (w/v) SDS, 0.03 % (w/v) ammonium persulfate (APS), 0.08 % (w/v) N,N,N’,N’-Tetramethyl ethylenediamine (TEMED)) and a 6 % stacking gel (6 % / 0.16 % (w/v) acrylamide/bis-acrylamide, 125 mM Tris-HCl pH 6.8, 0.1 % (w/v) SDS, 0.05 % (w/v) APS, 0.15 % (w/v) TEMED). The recipe for preparing one gel can be found in Tables 1 and 2, scale up as necessary.

2. Pre-stained protein ladder (available commercially).

3. Ponceau protein staining solution: 0.2 % (w/v) Ponceau S in 3 % (w/v) Trichloroacetic acid (TCA).

4. Nitrocellulose membrane (0.45 μm).

5. Wash buffer: 10 mM Tris-HCl pH 8.0, 1 mM EDTA pH 8.0, 150 mM NaCl, 0.1 % (v/v) Triton X-100.

6. Blocking buffer: 5 % (w/v) non-fat dry milk in wash buffer.

7. Primary antibodies against mitoribosome small and large subunit proteins and relevant assembly factors ( a list can be found in reference [29]).

8. Horseradish-peroxidase (HRP) conjugated secondary antibodies specific for the host species of primary antibodies.

9. Enhanced chemiluminescence (ECL) solution: ECL solution is made in house by combining equal volumes of solution 1 (100 mM Tris-HCl pH 8.5, 2.5 mM luminol, 400 μM p-coumaric acid) and solution 2 (100 mM Tris-HCl pH 8.5, 0.036 % (v/v) H₂O₂ (see Note 6). Store solutions 1 and 2 protected from light and at 4 °C, where they are stable for 1–2 months. The recipe for preparing these solutions can be found in Tables 3 and 4.

Table 1.

Recipe for 12 % acrylamide resolving gel.

Resolving gel – 12 % acrylamide
Reagent	[Stock]	[Final]	1x Volume (1 gel)
H2O	-	-	10.35 mL
Acrylamide/Bis-acrylamide (37.5:1)	30 %/0.8 %	12 %/0.32 %	9 mL
Tris-HCl pH 8.8	3 M	375 mM	2.8125 mL
SDS	10 %	0.1 %	225 μL
APS	10 %	0.03 %	75 μL
TEMED	~99 %	0.08 %	18.75 μL

Table 2.

Recipe for 6 % acrylamide stacking gel.

Stacking gel – 6 % acrylamide
Reagent	[Stock]	[Final]	1x Volume (1 gel)
H2O	-	-	5.28 mL
Acrylamide/Bis-acrylamide (37.5:1)	30 %/0.8 %	6 %/0.16 %	1.6 mL
Tris-HCl pH 6.8	1 M	125 mM	1 mL
SDS	10 %	0.1 %	80 μL
APS	10 %	0.05 %	40 μL
TEMED	~99 %	0.15 %	12 μL

Table 3.

Recipe to prepare chemiluminescence solution 1.

ECL Solution 1
Reagent	[Stock]	[Final]	1x Volume (50 mL)
H2O	-	-	44.28 mL
Tris-HCl pH 8.5	1 M	100 mM	5 mL
Luminol	250 mM	2.5 mM	500 μL
p-Coumaric acid	90 mM	400 μM	220 μL

Table 4.

Recipe to prepare chemiluminescence solution 2.

ECL Solution 2
Reagent	[Stock]	[Final]	1x Volume (50 mL)
H2O	-	-	44.94 mL
Tris-HCl pH 8.5	1 M	100 mM	5 mL
H2O2	30 %	0.036 %	60 μL

2.3. Mass Spectrometry Analysis of Assembly Intermediates Compositions and Clustering Analysis.

2.3.1. Methanol/Chloroform Protein Precipitation of Sucrose Gradient Fractions Containing Assembly Intermediates and Mitoribosomal Subunits

1. Methanol

2. Chloroform

3. High-speed vacuum concentrator

2.3.2. Clustering Analysis of Mass Spectrometry Data to Assign Hierarchical Assembly Clusters

1. RStudio software.

2. CSV file of proteomics results.

3. Methods

3.1. Generation of a Collection of Yeast Strains KO for Genes Coding for Mitoribosome Proteins and Assembly Factors

To generate a library of yeast mitoribosome knockout (KO) strains, we obtained a collection of deletion strains in which genes were replaced with a KanMX4 cassette conferring resistance to Geneticin. The strains are in a BY4741/4742 background and are available as part of the yeast knockout collection from Horizon Discovery Biosciences (Cambridge, United Kingdom). For this method, the KanMX4 cassette in each strain flanked by ~500 bp homologous to the MRP to be deleted was transformed in a diploid a/α W303I⁰ + VAR1^U + RNR1 [29]. Following sporulation and tetrads dissection, haploid deletion strains were obtained (Figure 1A) (see Note 7). Some strains, however, are not available as part of the collection and are created by replacing the entire ORF with a KanMX4 cassette or a HIS5 cassette.

Figure 1. Graphical outline of methods.

Schematic detailing the procedures for (A) generating a library of yeast deletion strains, (B) separating mitoribosome particles by sucrose gradient sedimentation. Figure modified from ref. [29], and (C) identifying assembly intermediates by tandem mass spectrometry. The figure was prepared with Adobe Illustrator and BioRender.

3.1.1. Amplification of Gene Targeted KanMX4 Deletion Cassettes

1. Design primers to amplify the KanMX4 cassette with an additional ~500 bp flanking the resistance marker that is homologous to the 5’- and 3’-UTRs of the corresponding mitoribosomal protein or assembly factor (Figure 1A). If the desired yeast null mutant is unavailable from the deletion strains in the Yeast Knockout Collection from Horizon Discovery Biosciences, see Note 8.

2. Extract genomic DNA following steps 3–13.

3. Culture yeast KO strain overnight in 10 mL of WO liquid media with 1x AHW.

4. Pellet 1 mL of confluent cells by centrifuging at 1,500 xg for 5 min at RT.

5. Wash cells by resuspending the pellet in 500 μL of sterile water and centrifuge again at 1,500 xg for 5 min at RT.

6. Resuspend cells in 150 μL of solution A and incubate at 37 °C for 1 h without shaking to digest the cell wall.

7. Add 20 μL of 10 % (w/v) SDS and vortex to mix.

8. Add 100 μL of 8 M ammonium acetate and vortex to mix.

9. Incubate at −20 °C for 15 min.

10. Centrifuge at 10,000 xg for 10 min at 4 °C.

11. Transfer 180 μL to a new microcentrifuge tube and add 120 μL of isopropanol. Vortex to mix and incubate for 5 min at RT to precipitate DNA.

12. Pellet the DNA by centrifuging at 10,000 xg for 15 min at 4 °C. Dispose of the supernatant and wash the pellet with 300 μL of 80 % ethanol.

13. Centrifuge at 10,000 xg for 1 min at 4 °C. Dispose of supernatant and air dry the pellet at RT. Resuspend the pellet in 30 μL of water.

14. Using genomic DNA extracted from the yeast KO strain of interest, amplify by PCR the gene-specific KO cassette. Isolate the correct product by separation on a 1 % (w/v) agarose gel in 1x TBE followed by DNA extraction with a commercially available kit.

15. Use the purified KO cassette for transformation into the diploid a/α W303I⁰ + VAR1^U + RNR1 strain.

3.1.2. Yeast Transformation

1. Inoculate a starter culture with the a/α W303I⁰ + VAR1^U + RNR1 strain in 10 mL of WO liquid media supplemented with the required auxotrophic markers (AHW) at a 1x working concentration and grow overnight at 30 °C, shaking at 200 xg.

2. Dilute starter culture in 10 mL of fresh WO liquid media supplemented with 1x AHW solution to an optical density at 600 nm (OD₆₀₀) = 0.1.

3. Continue to grow cells for 4–6 h until the culture reaches an OD₆₀₀ = 0.6–1.0.

4. Collect 1 mL of culture in a sterile microcentrifuge tube and pellet cells by centrifuging at 1,500 xg for 5 min at RT.

5. Wash cells by resuspending the cell pellet in 1 mL of sterile TEL and centrifuging again at 1,500 xg for 5 min at RT to pellet the cells.

6. Resuspend cells in 100 μL of sterile TEL. In a separate sterile tube, pre-mix 1–10 μg of the purified KO cassette from step 2 in section 3.1.1 with 10 μL of 10 mg/mL denatured salmon sperm carrier DNA (see Note 9). Add mixture to the cells and incubate for 30 min at RT without shaking.

7. Add 700 μL of sterile PEG-TEL and gently mix by pipetting. Incubate for 30 min at RT without shaking.

8. Heat shock at 42 °C for 10–15 min and transfer cells to ice (4 ^oC) for 2 min.

9. Centrifuge at 1,500 xg for 5 min at RT to pellet cells and resuspend in 1 mL of YPD liquid media. Outgrow cells for 30–60 min at 30 °C to allow recovery and increase transformation efficiency.

10. Pellet cells by centrifuging at 1,500 xg for 5 min at RT. Wash cell pellet by resuspending in 1 mL of sterile water and centrifuge again at 1,500 xg for 5 min at RT.

11. Resuspend the cell pellet in 100 μL of sterile water and spread on a WO solid media plate containing the selective resistance marker geneticin at final 500 μg/ml and the auxotrophic requirements (AHW) (see Note 10).

12. Incubate plates at 30 °C; colonies should appear in 4–5 days. Pick colonies and test them by genotyping.

3.1.3. Genomic DNA Extraction and Validation of KanMX4 Cassette Integration

1. Extract genomic DNA as explained in section 3.1.1 (steps 2–13).

2. Validate correct KanMX4 cassette integration by PCR using primers that amplify regions spanning the KanMX4 cassette and the 5’- and 3’-UTRs to ensure deletion in the correct genetic locus (Figure 1A).

3. Assess that the deletions strains retain mtDNA (see Note 7).

3.2. Sucrose Gradient Analysis to Establish the Mitoribosome Profile and Detect Assembly Intermediates.

The overall approach is depicted in Figure 1B.

3.2.1. Culture of Yeast Cells

S. cerevisiae can utilize a variety of different carbon sources to transduce energy and fuel growth. Fermentable carbon sources, such as glucose and galactose, drive energy production through glycolysis-based metabolism. Alternatively, cells grown with non-fermentable carbon sources like glycerol and ethanol adapt to an aerobic-based metabolism, generating ATP through cellular respiration. Yeast cells are highly sensitive to changes in metabolism and regulate mitochondria quantity, morphology, and proteome depending on the available carbon source [35–37]. When available, yeast preferentially utilize glucose to generate energy via glycolysis, regardless of the available oxygen [35, 38]. Consequently, under high glucose conditions, the expression of genes encoding enzymes of the TCA cycle and many respiratory complex proteins is downregulated, resulting in impaired OXPHOS complex assembly and activity [39–41]. This phenomenon is known as “glucose repression” and inhibits the use of other carbon sources and respiration-based metabolism via signaling, transcriptional, and post-translational regulation [35, 38, 41]. Many studies have proposed that mitoribosome activity is tightly regulated to coordinate the synthesis of mtDNA-encoded proteins in response to OXPHOS complex assembly demands [42–44]. Interestingly, a recent study suggests that mitochondrial translation capacity is maintained under high glucose conditions and may be uncoupled from the repressive effects on OXPHOS assembly and activity [45]. Galactose however is a non-repressive fermentable carbon source. For this reason, cells are grown in WO-Gal to isolate mitochondria in this protocol (see Note 11).

1. Yeast strains should be spread on WO-Gal-AHW solid media agar plates from a rho+ glycerol stock stored at −80 °C. Incubate plate at 30 °C and use within 2–3 days to inoculate cultures.

2. For each yeast strain, inoculate 4 separate flasks at a starting OD₆₀₀ = 0.01 in 100 mL of WO-Gal-AHW medium (4× 100 mL). Grow starter cultures for 24–48 h at 30 °C with shaking, taking periodic OD₆₀₀ measurements to estimate doubling time (see Note 12).

3. Per flask, dilute starter culture into 1 L of WO-Gal-AHW (4× 1 L per strain) and grow at 30 °C with shaking until OD₆₀₀ = 1.5–2.0 (see Note 13). Be careful not to exceed 20 generations of growth (see Note 14). Proceed directly to mitochondrial isolation.

3.2.2. Isolation of Mitochondria

Isolation of mitochondria as described in this protocol yields a highly enriched mitochondrial preparation with intact-membrane organelles. A high-quality mitochondrial isolate is essential for meaningful downstream proteomics analysis. This method is slightly modified from Horn et al. (2008) [46] to increase the removal of common non-mitochondrial co-contaminants such as the ER and Golgi. In brief, this protocol is based on the homogenization of spheroplasts followed by the isolation of mitochondria by differential centrifugation. Mitochondria prepared using this technique have intact membranes and can be used for a variety of downstream in organello analyses such as oxygen rate consumption, protein synthesis and transport, and transcription.

1. Harvest cells in polypropylene centrifuge bottles by centrifuging at 900 xg for 5 min. Wash cells once with sterile RT water.

2. Resuspend cells in Tris-DTT buffer at a ratio of 1 mL per 2 g of cells and incubate at 30 °C for 10 min with gentle shaking (see Notes 15 and 16).

3. Pellet the cells by centrifugation at 2,200 xg for 6 min and wash cells once with 1.2 M sorbitol using 150 mL per 10 g of cells (see Note 17).

4. Resuspend cells in digestion buffer using 1 mL for every 0.15 g of cells (see Note 18). Incubate at 30 °C for 30 min with gentle shaking. At the end of the incubation period immediately place bottles containing spheroplasts on ice to stop the digestion reaction (see Note 19). All subsequent steps must be done at 4 °C.

5. Pellet spheroplasts by centrifugation at 1,250 xg for 5 min at 4 °C. Wash once with ice-cold 1.2 M sorbitol.

6. Resuspend washed spheroplasts with ice-cold homogenization buffer using 1 mL for every 0.15 g of cells (see Note 20). Transfer to a pre-chilled 50 mL glass dounce tissue homogenizer fitted to a loose Teflon pestle and homogenize with 6–8 strokes (see Note 21).

7. Centrifuge homogenate at 1,700 xg for 6 min at 4 °C. Repeat this step on the supernatant in a new centrifuge bottle.

8. Centrifuge the supernatant at 14,000 xg for 6 min at 4 °C to pellet mitochondria.

9. Resuspend pelleted mitochondria in 10 mL of SH buffer (see Note 22). Centrifuge at 2,200 xg for 5 min at 4 °C.

10. Centrifuge the supernatant containing mitochondria at 14,000 xg for 6 min at 4 °C. Wash mitochondrial pellet with 10 mL of ice-cold SH buffer to remove additional contaminants (see Note 23). Resuspend mitochondria in 200–600 μL of cold SH buffer, depending on pellet size (see Note 24).

11. Determine protein concentration using the desired method and adjust to 10 mg/mL. Prepare 2–4 mg-aliquots that will be used for mitoribosome extraction and sucrose gradient sedimentation in the subsequent steps. Use the remainder of the mitochondria to prepare samples for immunoblot analysis (see Note 25). Flash freeze samples in liquid nitrogen and store at −80 °C until use (see Note 26).

3.2.3. Preparation of a 10–30 % Linear Sucrose Gradient

Density gradients are a valuable separation technique that can be used to isolate particles in a mixture of biomolecules or further purify organelles. The approach described here uses rate zonal separation in which particles sediment based on mass and size (not only density) and all particles will eventually accumulate at the bottom of the tube if centrifuged long enough. We describe here the preparation of a continuous 10–30 % sucrose linear gradient prepared by using a titled tube rotation approach, the Biocomp Instruments Gradient Master. These gradients have been shown to successfully separate mitoribosome subpopulations representing assembly intermediates in addition to fully assembled subunits and monosomes from a mitochondrial lysate [29, 47].

1. Prepare both 10 % and 30 % sucrose gradient buffers according to the recipe in Table 5 (see Notes 27, and 28). Place solutions on ice to cool.

2. Begin preparing the gradient by placing the 10 mL ultracentrifuge tube in the marker block. Mark the upper edge with a fine tip marker (see Note 29). This will indicate the “stopping point” when filling with sucrose gradient solution. Place marked ultracentrifuge tube in the MagnaBase tube holder.

3. Fill the ultracentrifuge tube with the 10 % sucrose solution until the volume is slightly above the marked stopping point (~2 mm) (see Note 30).

4. Using the provided layering cannula attached to a syringe, insert the cannula tip to the bottom of the tube and slowly fill with the 30 % sucrose solution. As the solution is added from the bottom of the tube the lighter 10 % sucrose layer will be pushed toward the top of the tube. A clear interface between the two layers should be visible. Fill solution until just above the marked stopping point. Once added, swiftly and carefully remove the cannula.

5. Place 4 mm tube cap on ultracentrifuge tube (see Note 31).

6. Use a leveling tool to ensure that the magnetic plate on the gradient master is perfectly level.

7. Carefully place the tube holder with sucrose gradients onto the center of the gradient master plate (see Note 32). Ensure that tubes are balanced.

8. Use gradient master to make a 10–30 % linear sucrose gradient: Using the gradient master manual interface select ‘Grad’ ‘List’ ‘SW55’ ‘10–30 %’ ‘Run’ (see Note 33).

9. Gently remove gradients from the tube holder and keep them at 4 °C until used in the next step (see Note 34).

Table 5.

Recipe for 10 % and 30 % sucrose solutions to prepare sucrose gradients. The recipe indicates the volumes required for two samples. Scale-up as necessary.

Gradient Buffer
			10 % sucrose	30 % sucrose
Reagent	[Stock]	[Final]	2x Volume (5 mL)	2x Volume (5 mL)
Tris pH 7.4	1 M	10 mM	50 μL	50 μL
NH4Cl	1 M	100 mM	500 μL	500 μL
MgCl2	1 M	10 mM	50 μL	50 μL
PMSF	100 mM	1 mM	50 μL	50 μL
Digitonin (high purity)	10 %	0.05 %	25 μL	25 μL
Yeast protease inhibitor cocktail	100x	1x	50 μL	50 μL
RNase inhibitor	40 U/μL	0.1 U/μL	12.5 μL	12.5 μL
Sucrose	50 %	10 %/30 %	1000 μL	3000 μL
H2O	-	-	3262.5 μL	1262.5 μL

3.2.4. Mitoribosome Extraction and Sedimentation Analysis

To ensure mitoribosomes and any possible assembly complexes remain intact, mitoribosomes are extracted with the mild non-ionic detergent digitonin. A crucial aspect in maintaining the association between the small and large subunits in the mitoribosome is the ratio of monovalent to divalent cations in both the extraction and sucrose gradient buffers [43, 48]. A 10:1 ratio of monovalent to divalent ions using 100 mM NH₄Cl and 10 mM MgCl₂ is described here. Additionally, the use of an RNase inhibitor to prevent the degradation of rRNA and the bound mRNA may help to stabilize the monosome. Take care to avoid the use of EDTA and other ion chelating agents as this will disrupt the interaction between subunits. An example of the dissociation caused by EDTA can be observed in Figure 1B. Sample fractionation in this protocol is performed with a straight needle, however, a side hole needle or piston fractionator fitted with a trumpet needle will provide increased resolution.

1. Remove 2–4 mg aliquot of isolated mitochondria from −80 °C and thaw on ice (see Note 35). Use immediately upon thawing.

2. Centrifuge at 10,000 xg for 10 min at 4 °C to pellet mitochondria.

3. Remove the supernatant and resuspend the mitochondrial pellet in 400 μL of ice-cold freshly prepared extraction buffer. Incubate on ice for 10 min.

4. Centrifuge at 24,000 xg for 15 min at 4 °C. Collect supernatant containing mitoribosomal and soluble mitochondrial proteins as mitochondrial extract (see Note 36).

5. Carefully pipette the remaining supernatant (~360 μL) onto the top of the prepared 10–30 % sucrose gradient (see Note 37). Add slowly to ensure that the sample does not mix with the upper sucrose layer.

6. Gently place sucrose gradient into the swinging-bucket rotor. Cap tightly and ensure that the rotor is balanced. Ultracentrifuge at 200,000 xg for 3 h: 10 min at 4 °C (see Note 38). Once finished, very carefully remove the gradient from the rotor. Avoid any disturbances at this point as this will disrupt the sedimentation profile of protein complexes in the gradient. Keep gradient cold until proceeding to fractionation.

7. Fractionate sucrose gradient by piercing the bottom of the ultracentrifuge tube with a standard straight beveled needle. Collect 14 equal volume fractions drop-wise in microcentrifuge tubes (see Note 39). Immediately place all fractions on ice.

8. Immediately prepare fraction samples for immunoblotting by combining 75 μL of each fraction with 25 μL of 4x LB (see Note 40). Store the remainder of each fraction at −80 °C until used for protein precipitation and subsequent mass spectrometry analysis.

3.2.5. SDS-PAGE and Immunoblot Analysis of Sedimented Mitoribosomal Particles

To identify assembly intermediates for downstream proteomics, the mitoribosome sedimentation profile and distribution of subassembly particles can be analyzed by immunoblot. Comparison of assembly intermediates with the fully assembled subunit in the WT may suggest if the protein deleted acts in an early or late stage of the assembly process. For example, if an early assembly protein is deleted, intermediates will accumulate in the lightest fractions of the gradient and be significantly smaller than the WT particle. Alternatively, assembly intermediates in late assembly mutants will be larger and have a sedimentation profile like WT. Figure 1B shows the distribution of subassemblies that accumulate in the early, intermediate, and late stages of mtLSU assembly. Use as many different tested antibodies possible to ensure visualization of intermediates by immunoblot. Alternatively, although not described here, gradient fractions could be screened by northern blotting or RT-qPCR for the presence of mitochondrial rRNA. Fractions identified to harbor intermediates will be further analyzed by mass spectrometry.

1. In a 12 % SDS-PAGE gel made with a 16-lane comb, load from left to right a pre-stained protein ladder, 10 μL of the “total extract” from step 4 in section 3.2.4, and 30 μL each of the fractions 1 through 14 from step 8 in section 3.2.4.

2. Begin electrophoresis at 100 V and allow samples to migrate through the stacking portion of the gel (~30 min) (see Note 41). Once the Laemmli sample buffer dye front enters the resolving gel increase the voltage to 140 V and run until the dye front reaches the bottom of the gel (~4 h).

3. Transfer proteins to a nitrocellulose membrane by semi-dry transfer at 250 mA for 5 h. Alternatively, a wet transfer can also be used.

4. After transfer, wash the membrane with water for ~15 min. To check for protein degradation and assess the distribution of protein across fractions, incubate the membrane with ponceau staining solution with mild shaking for ~15 min (see Note 42).

5. Using the protein ladder as a reference guide, cut the membrane in strips such that each piece corresponds with the molecular weight of the mtSSU or mtLSU protein to be probed against by the predetermined antibody.

6. Rinse membranes in wash buffer with mild shaking for 5–10 min or until the ponceau stain has been washed out.

7. Block membranes by incubating in blocking buffer for 30 min with mild shaking. Alternatively, membranes can be blocked overnight at 4 °C.

8. Incubate membranes with respective primary antibodies for 1 h at RT with mild shaking (see Notes 43-45).

9. Rinse membranes by incubating in wash buffer with mild shaking for 10 min. Perform three washes in total.

10. Incubate membranes in secondary antibody for 1 h at RT with mild shaking (see Note 46).

11. Rinse membranes again by performing three washes in wash buffer with mild shaking for 10 min each.

12. Develop membranes by incubating them in ECL solution and subsequently exposing them to autoradiography film. Identify fractions corresponding to any assembly intermediates in addition to fractions representing the mtSSU, mtLSU, and monosome. These fractions will be used for protein precipitation and quantitative mass spectrometry.

3.3. Mass Spectrometry Analysis of Assembly Intermediates Compositions and Clustering Analysis

The overall approach is depicted in Figure 1C.

3.3.1. Methanol/Chloroform Protein Precipitation of Sucrose Gradient Fractions Containing Assembly Intermediates and Mitoribosomal Subunits

An essential step in ensuring accurate identification of assembly intermediates and collection of high-quality data is to prepare samples for mass spectrometry analysis by precipitating protein from sucrose gradient fractions. Methanol/chloroform extraction allows for the elimination of lipids, detergents, nucleic acids, salts, and other contaminants that may otherwise interfere with ionization or data interpretation (Figure 1C).

1. Per 100 μL of sample add 400 μL of methanol and vortex well to mix (see Note 47).

2. Add 100 μL of chloroform and vortex well to mix (see Note 48).

3. Add 300 μL of water and vortex well to mix (see Note 49). The sample should become cloudy as phase separation begins to occur.

4. Centrifuge at 14,000 xg for 2 min.

5. Carefully remove the aqueous (top) phase with a pipette and discard. Proteins reside at the thin interphase between the organic and aqueous layers and are easily disturbed.

6. Add 400 μL of methanol and vortex well to mix (see Note 50).

7. Centrifuge at 14,000 xg for 3 min.

8. Remove as much solvent as possible by pipetting without disturbing the protein pellet.

9. Dry the pellet using a high-speed vacuum concentrator (see Note 51).

10. Send samples to a facility for LC-MS/MS (liquid chromatography-tandem mass spectrometry) analysis.

3.3.2. Quantitative Mass Spectrometry Analysis of Assembly Intermediates and Data Clustering Analysis to Determine Hierarchical Mitoribosome Protein Assembly

Analyzing samples by nano LC-MS/MS allows for highly sensitive quantification and accurate detection of proteins in a mixture. This method enables proper identification and classification of assembly intermediates as well as any previously unknown assembly factors, which may help in defining a de novo assembly pathway. Liquid chromatography (LC) coupled to mass spectrometry (MS) allows for the physical separation of components in a mixture prior to mass analysis. Tandem mass spectrometry (MS/MS) involves using two mass analyzers and further fragmentation and separation of precursor ions to provide more sensitive detection of proteins and other biomolecules. In short, the process of LC-MS/MS begins with the enzymatic cleavage of proteins into small peptides, typically by trypsin digestion. Peptides are then separated by liquid chromatography and eluted based on their affinity for the stationary phase and the solvent used. Following this, peptides are passed to the mass spectrometer and ionized by electrospray or another equivalent ionization method. The mass analyzer then separates the precursor ions based on their mass-to-charge ratio (m/z). These “parent” ions are then further fragmented by collision-induced dissociation (CID). The ion fragments are separated by the second mass analyzer before finally being measured by the detector. The detector will produce a mass spectrum for each ion. An increasingly popular and cost-efficient method for MS-based quantitation and peptide identification is the use of isobaric stable isotope labeling. For example, tandem mass tag (TMT) mass spectrometry yields robust and reproducible protein quantitation while enabling sample multiplexing up to 16 plex. Using data analysis platforms such as Proteome Discoverer or MaxQuant, the fragmentation spectra are referenced against a protein sequence database by search algorithms including MASCOT, SEQUEST, X!TANDEM, Andromeda, or any other commonly used database search engine. All bioinformatics and statistical analysis should be done using the TMT reporter intensity. However, in this chapter, we present the use of values from the MASCOT algorithm and ion search engine. MASCOT is a powerful tool that provides probabilistic scoring and integrates peptide mass fingerprints, protein sequence queries, and tandem MS ion searches. Based on this, a protein score or “MASCOT score” is distilled in which the value represents the combined score of each ion fragment spectra matching peptide sequences within the same protein. This value, however, is a probability, and for increased sensitivity, TMT reporter intensity should be used in place of MASCOT scores. LC-MS/MS analysis should be performed in at least biological triplicate. Differential regulation across all groups is tested by ANOVA, and Tukey’s HSD test can be used to make comparisons between pairs. MASCOT scores can be used for downstream hierarchical clustering analysis to determine assembly modules as described below.

1. In preparation for clustering analysis and to make comparisons between strains, MASCOT scores for each protein should be normalized to WT and Log₂ transformed $(Lo{g}_2\frac{mutan{t}_{MASCOT}}{W{T}_{MASCOT}})$. Log transformation reduces heteroscedasticity and results in a more normal distribution of the data which in turn increases the strength of statistical tests.

2. To facilitate cluster analysis, arrange data in a CSV file such that rows represent observations (mutant strains) and columns indicate variables (MASCOT score for each protein).

3. Import the dataset into RStudio and load the packages “gplots” and “RColorBrewer” (see Note 52).

4. Use the heatmap.2 function to generate a heatmap of the dataset. The function will automatically perform hierarchical clustering for both rows and columns and reorder the corresponding dendrograms based on the mean. This will yield vertical clusters of proteins that maintain similar stabilities across mutant strains. Vertical clustering of these stability profiles yields distinct mitoribosome structural modules (Figure 2). One such example from the yeast mtLSU is the central protuberance, in which proteins composing this structural module are detected at similar levels across all mutant strains (Figure 2). Horizontal clustering generates assembly clusters representing mutant strains with similar mitoribosome protein subassembly profiles (Figure 2). The similarity of the mitoribosome protein composition for mutant strains in each assembly cluster indicates that the deleted proteins act at the same or similar stage of assembly.

5. Assembly clusters can then be classified as either early, intermediate, or late incorporation modules based on the relative stability and detection of all other mitoribosomal proteins. For example, mutant strains in the earliest assembled cluster will have subassemblies in which most proteins are decreased or undetectable, as the early assembly proteins serve as a scaffold for subsequent clusters. On the contrary, mutant strains of a late assembly cluster will have subassemblies in which most proteins are present at WT levels. In the case of the yeast mtLSU the first assembled cluster, depicted in red, correlates with the most underrepresentation of mitoribosomal proteins and has extensive contact with the ribosomal RNA (Figure 2). Whereas subassemblies of proteins in the last assembled cluster, shown in orange, are characterized by either WT levels or an increase in most mitoribosomal proteins (Figure 2). Constituents of late assembly clusters typically map to the surface of the mitoribosome (Figure 2).

6. Additional insight from phenotypic experiments such as ribosomal RNA levels, cellular respiration, and protein synthesis capacity combined with structural comparison to bacterial counterparts allows for the construction of a more granular pathway and a hierarchical ranking of clusters.

Figure 2. Clustering analysis and elucidation of hierarchical mitoribosome large subunit assembly pathway.

(A) Heatmap of relative protein stability (vertical axis) in each yeast deletion strain (horizontal axis) as determined by mass spectrometry of subassembly containing sucrose gradient fractions. Structural modules are clustered on the vertical axis and their structures are mapped to the mtLSU with the 21S rRNA shown in grey. Assembly modules are clustered on the horizontal axis with their representative cryo-EM structures shown below. Each cluster is color coded to match the corresponding dendrogram. Figure modified from ref. [29]. (B) Assembly pathway mapping clusters defined in panel A to the mtLSU. Clusters are denoted either early (red), intermediate (cyan, yellow, blue), or late (purple, orange). The 21S rRNA is shown in grey. All structures are prepared using PDB: 3J9M with proteins shown as surface.

Table 6.

Primer pair used to generate library of yeast KO strains.

Description	Sequence (5’ – 3’)
Amplification of kanMX4 cassette	F: CGTACGCTGCAGGTCGAC R: ATCGATGAATTCGAGCTCG

Table 7.

Recipe to prepare solution for extraction of mitoribosomes. This solution can be made at the same time as preparing the sucrose gradient buffers. Ensure the digitonin used is high-purity and is prepared the same day as the experiment.

Extraction Buffer
Reagent	[Stock]	[Final]	1x Volume (500 μL)
Tris pH 7.4	1 M	10 mM	5 μL
NH4Cl	1 M	100 mM	50 μL
MgCl2	1 M	10 mM	5 μL
PMSF	100 mM	1 mM	5 μL
DTT	100 mM	0.25 mM	1.25 μL
Digitonin (high purity)	10 %	0.6 %	30 μL
Yeast protease inhibitor cocktail	100x	1x	5 μL
RNase inhibitor	40 U/μL	0.1 U/μL	1.25 μL
H2O	-	-	397.5 μL