Stable Isotope Labeling with Amino acids in Cell culture, known as SILAC, is a metabolic labeling approach used in quantitative mass spectrometry-based proteomics. This technique enables precise comparison of protein abundance between two or more distinct cell populations. Researchers employ SILAC to explore various cellular processes, including responses to drug treatments, signaling pathways, or protein-protein interactions.
The Principle of SILAC Labeling
SILAC operates by metabolically incorporating non-radioactive, “heavy” stable-isotope-containing amino acids into living cells’ proteome. Cells are grown in specialized media where naturally occurring “light” amino acids are replaced with their isotopically enriched counterparts. Over several cell divisions, these labeled amino acids integrate into all newly synthesized proteins. This creates two cell populations: one with “light” proteins and another with “heavy,” labeled proteins.
Specific amino acids, L-Arginine and L-Lysine, are chosen for this labeling. These are essential amino acids, which cells cannot synthesize. The enzyme trypsin, commonly used for protein digestion in proteomics, specifically cleaves proteins at the C-terminal side of arginine and lysine residues. This ensures nearly every peptide from digestion contains a labeled arginine or lysine, allowing comprehensive proteome labeling. The isotopic difference, often involving ¹³C or ¹⁵N atoms, provides a distinct mass tag detectable by a mass spectrometer.
Essential Materials and Cell Culture Preparation
Initiating a SILAC experiment requires specialized materials for accurate labeling. Key materials include cell culture media deficient in L-lysine and L-arginine, forcing cells to incorporate supplied amino acids. Dialyzed fetal bovine serum (dFBS) is also necessary, as it removes naturally occurring “light” amino acids that would compete with isotopic labels.
Along with deficient media and dFBS, researchers prepare “light” and “heavy” isotopic amino acids. Light amino acids are unlabeled L-lysine and L-arginine. For “heavy” labels, researchers commonly use L-lysine with six ¹³C atoms (Lys6) and L-arginine with six ¹³C atoms (Arg6), or variations involving ¹⁵N isotopes. Before labeling, cells are seeded and grown to ensure they are healthy and actively dividing.
The Core SILAC Experimental Workflow
The SILAC workflow begins with cell adaptation. One cell population is cultured in “light” media containing unlabeled amino acids, while a parallel population is grown in “heavy” media containing isotopically labeled L-lysine and L-arginine. Cells are maintained in these media for typically five to six cell divisions. This ensures nearly all intracellular proteins incorporate the labeled amino acids, achieving high labeling efficiency.
After adaptation, a sample from the “heavy” population is often analyzed by mass spectrometry to confirm label incorporation. This ensures over 95% of proteins incorporate stable isotopes before the main experiment. Once sufficient labeling is verified, the specific experimental treatment can be applied. For instance, one cell population might receive a drug treatment, while the other serves as an untreated control, allowing for direct comparison of protein changes.
Following treatment, both cell populations are harvested. Accurate cell counting is performed for each population to ensure precise mixing ratios. The “light” and “heavy” cell populations are then mixed, typically at a 1:1 ratio. Combining samples at this early stage is an important aspect of SILAC, as it minimizes experimental variability and technical errors during downstream processing.
Sample Preparation for Mass Spectrometry
After the “light” and “heavy” cell populations are harvested and mixed, proteins are prepared for mass spectrometry analysis. The mixed cell pellet undergoes lysis to extract proteins. Lysis buffers with detergents and protease inhibitors ensure complete protein solubilization and prevent degradation. The resulting protein solution represents a combined sample of both experimental conditions.
Following protein extraction, proteins are subjected to enzymatic digestion, most commonly using trypsin. Trypsin cleaves proteins into smaller fragments called peptides at specific sites, primarily after lysine and arginine residues. This digestion is performed under controlled conditions for consistent and complete cleavage, important for mass spectrometry analysis. The resulting peptide mixture is then ready for purification.
Purification involves sample cleanup and desalting to remove detergents, salts, and other interfering compounds that can negatively impact mass spectrometry performance. Techniques such as C18 StageTips or ZipTips are frequently used. These methods selectively bind peptides while allowing contaminants to pass through, resulting in a cleaner, more concentrated peptide sample. The purified peptides are then ready for introduction into the mass spectrometer.
Data Acquisition and Interpretation
The prepared peptide mixture, containing both “light” and “heavy” versions of the same peptides, is analyzed using Liquid Chromatography-Mass Spectrometry (LC-MS/MS). Liquid chromatography separates the complex mixture of peptides based on their physicochemical properties, while the mass spectrometer detects and measures these separated peptides. The mass spectrometer identifies pairs of chemically identical peptides that differ only by mass, corresponding to “light” and “heavy” versions.
For each identified peptide pair, the mass spectrometer measures the signal intensity of both the “light” and “heavy” forms. The ratio of these signal intensities directly reflects the relative protein abundance in control versus treated samples. For example, a higher signal for the “heavy” peptide compared to its “light” counterpart indicates an increased abundance of that specific protein in the treated cells. Specialized software, such as MaxQuant, automates peptide identification, quantification, and protein abundance determination from raw mass spectrometry data.