This work was supported by the National Institutes of Health (NIH) and the Intramural Research Program (IRP), National Institute on Aging (NIA). N.B. was supported by Longevity Impetus Grants, and the Office of Dietary Supplements (ODS) Scholars Program. Figure 1 was created with BioRender.com.

The authors have no disclosures.

pSILAC is a powerful technique that enables the global quantification of protein turnover rates across multiple cellular conditions. This paper details the use of pSILAC to compare global protein half-lives between senescent and quiescent cells, including instructions for the preparation of senescent and quiescent cells, SILAC labeling and harvesting, and ultimately analysis using DDA mass spectrometry. Additionally, a two-step test is described for validation of the senescence phenotype using SA-&#946;Gal and RT-qPCR analysis of a panel of mRNAs encoding senescence-associated proteins. In addition to validation of senescence with the two approaches described, a third validation of senescence can be performed following mass spectrometry analysis by looking for changes in known senescence markers between senescent and quiescent cells at the proteomic level. Senescence-associated proteins that are expected to be elevated include p16, p21, and BCL2, among others, described elsewhere44,45. In the protocol described above, ionizing radiation was used for the induction of senescence and serum starvation for quiescent cells. For the induction of senescence, there are multiple options available and there is substantial heterogeneity among them41,42,46. Currently, there is no senescent method that is considered the &#34;most physiological&#34;, so the choice of senescent inducer is largely based on the context of the experiment. However, experiments with the goal of stating a general phenomenon about senescence are recommended to use at least two different senescent inducers. Discussing the range of senescence paradigms is beyond the scope of this paper, but some common methods to induce senescence include triggering DNA damage (IR, doxorubicin, replicative exhaustion), expressing oncogenic proteins (HRAS, BRAF), and disrupting mitochondria function2.
In addition to the choice of senescent inducer, the choice of control cells is an equally important consideration. Senescent cells are, by definition, under indefinite growth arrest, so a comparison to other growth-arrested cells are often chosen. For pSILAC, cell cycle arrested cells are generally preferable because they do not replicate and are thus easier to use for protein half-life calculations47. However, since cultured cells will often retain some dividing cells, it is important that the methods used to induce cell cycle arrest produce as homogenous a response as possible to minimize error from cells that are still proliferating. To calculate protein degradation rates for cycling cells using pSILAC requires additional calculations to compensate for the rate at which protein is diluted into daughter cells27. However, quiescent growth arrest itself is not without complications. There are two general methods for cell cycle arrest: serum deprivation and contact inhibition48. Not all cells can be made quiescent through contact inhibition, although some fibroblasts have been shown to demonstrate quiescence after several days of culturing49. This method used serum deprivation because it is more commonly used for comparisons of senescent cells, though it requires for the senescent cell to be similarly serum-deprived for accurate comparisons. Serum activates the mTOR complex, and thus serum deprivation has several downstream effects on the cell in addition to cell cycle arrest50. Notably, senescent cells have been shown to display a reduced SASP upon serum deprivation or mTOR inhibition51,52.
Another important point to consider in pSILAC is how many time points to test. This protocol collected cells at a single time point (3 days of light or heavy labeling), which substantially simplifies the resulting analysis. The choice of time point should be based on the objective of the experiment. For global analysis, 3 days is expected to capture a majority of proteins, though half-lives for short-lived proteins that completely turnover within 3 days (all light signal is lost) cannot be measured at this time point. Conversely, long-lived proteins with very little turnover in 3 days are also difficult to quantify and often appear as having extremely large half-lives (in the order of weeks) that are typically just a consequence of very little heavy signal accumulation. Due to the non-linear relationship in the ratio of heavy and light peptide signals versus the percentage of newly synthesized protein at shorter and longer time points, the quantitation of half-lives could be improved by adding additional labeling time points. For relative comparisons between two cell states, as in this protocol, an approximate half-life may be sufficient, but additional timepoints can be used to improve quantitative accuracy.
This protocol describes how to perform an untargeted DDA-based analysis of protein turnover. However, the protein turnover calculations can be applied generally to any acquisition scheme that is able to derive the relative abundance of heavy and light peptide pairs. For example, MS2-based methods such as data-independent acquisition (DIA/SWATH) can also be applied for the calculation of turnover rates successfully53. Additionally, instrumentation and software pipelines other than those described in this protocol can be used to perform DDA analysis, protein identification, and protein quantification. When using protein quantification software platforms such as Skyline to extract peptide peak areas, it is advisable to manually inspect extracted ion chromatograms in the document workspace, identify peaks that were erroneously integrated and non-quantitative peaks, and curate the document accordingly. An extensive collection of tutorials is available online for Skyline (skyline.ms).
pSILAC represents one of the most ideal methods for global quantification of protein half-lives in cultured cells due to superior multiplexing (proteome coverage) and throughput. While pSILAC does not provide direct rates of synthesis or degradation, since the change in light and heavy signal is due to a confluence of factors, pSILAC is highly useful for comparisons between conditions and different cell types. Low-throughput methods often fall into two types: 1) treatment of cells with cycloheximide to block protein synthesis and harvest at time intervals after addition to monitor decay, or 2) treatment of cells with an inhibitor of protein decay and harvest at time intervals after addition to monitor the accumulation of protein, thus inferring protein decay rates. The limitation of both methods is that such treatments will inevitably cause substantial changes to cellular physiology. In contrast, pSILAC requires no substantial intervention and theoretically has no detectable effects on cellular physiology since isotopic amino acids differ by only a single neutron from their non-isotopic counterparts. Thus, the method described here for pSILAC represents a simple protocol for global measurement of the most physiological protein half-lives in non-dividing cells.
Alterations in protein turnover have a close relationship to aging, age-related diseases, neurodegeneration, and longevity54,55. This protocol describes a method to interrogate these relationships by using stable isotope labeling of amino acids in cell culture to measure protein turnover rates in senescence cells. However, numerous analogous methods exist to perform studies in the context of aging and neurodegeneration in vivo in whole organisms such as mice. Indeed, these studies have emphasized the importance of measuring protein turnover rates in the context of age-related diseases56,57,58,59.
In this study, ribosomal proteins and proteins residing in the endoplasmic reticulum&#160;stood out as two categories of proteins with decreased and increased half-lives in senescent cells, respectively. While further analysis of steady-state levels is required for definitive conclusions, these results further suggest that senescent cells may uniquely regulate translation through decreased half-lives of ribosomal proteins. Going forward, applying stable isotope labeling approaches to study the relationship between cellular senescence and neurodegeneration in vivo in mouse models will be a promising extension of the isotope labeling approach described by this protocol.

Senescence was first identified as a state of indefinite growth arrest exhibited by cultured primary cells after reaching replicative exhaustion1. It has since been shown that senescence can arise in response to numerous cellular insults, including genotoxic, mitochondrial, and oncogenic stresses, among others2. While senescence has several physiologically important roles, such as tumor suppression and wound healing, the accumulation of senescent cells during aging is associated with a host of deleterious effects on health3, including several neurodegenerative conditions4,5,6. Cellular senescence occurs in multiple brain cell types, including neurons7,8,9,10, astrocytes11, microglia12, and oligodendrocyte precursors13, and contributes to neurodegeneration and cognitive dysfunction. Amyloid beta oligomers, one of the hallmarks of Alzheimer&#39;s disease14, have been shown to accelerate neuronal senescence13,15,16. An increased prevalence of senescent cells has also been associated with Parkinson&#39;s disease17, especially arising from environmental stressors11,18. Importantly, the selective elimination of senescent cells in pre-clinical models extends lifespan and mitigates a multitude of age-related diseases3,5,12 and improves cognitive deficits8,11,12,13. Senescent cells have thus emerged as promising therapeutic targets for the treatment of many age-related conditions.
Much of the detrimental effect of senescent cells is caused by the senescence-associated secretory phenotype (SASP), a complex mixture of bioactive molecules secreted by senescent cells that can cause local inflammation, angiogenesis, destruction of the extracellular matrix, and propagation of senescence in surrounding tissue19,20,21. The SASP also represents an interesting biological phenomenon of senescence because it requires a considerable transcriptional and translational effort during a state of cell cycle arrest. In fact, senescent cells have been shown to exhibit decreases in ribosome biogenesis22,23,24 that should reduce protein synthesis. Instead, senescent cells robustly translate some proteins, particularly SASP factors, and influence the metabolism of surrounding tissue25. Thus, there is considerable interest in understanding how senescent cells undergoing a permanent cell cycle arrest continue to maintain protein homeostasis while at the same time robustly express SASP factors and other select proteins.
This method describes how to use mass spectrometry and pulsed-stable isotope labeling by amino acids in cell culture (pSILAC) to globally measure the half-lives of proteins in senescent cells at a proteome-wide scale. In traditional SILAC, cultured cells are completely metabolically labeled with heavy and light non-radioactive isotopes of amino acids for downstream analysis of protein abundance. This method has been previously applied to assess abundance changes comprehensively and quantitatively in the SASP of cultured fibroblasts26. In pSILAC, cells are similarly metabolically labeled with a pulse of heavy isotope that follows pre-labeling with light isotope, and then harvested at one or more time intervals. The rates of incorporation of heavy isotope in reference to pre-existing light isotope are then used to calculate relative protein turnover rates. Generally, isotopes of arginine and lysine are used because trypsin cleaves at those residues; thus, all peptides from standard digestion will potentially contain the heavy label. Pairs of peptides that differ only by the presence or absence of heavy lysine or arginine are chemically identical and can be differentiated and quantified by a mass spectrometer. Following mass spectrometry analysis, peptides can be identified as newly synthesized or pre-existing based on the presence or absence of the isotopic label in the resulting peptide identifications. Protein turnover rates can then be determined by fitting the ratio of heavy (13C-15N) over light (12C-14N) peptides for a given protein to kinetic models for exponential growth or decay27,28. pSILAC has been used in several comparisons of protein turnover rates29,30,31,32 and is currently the most comprehensive and high-throughput method for the measurement of protein half-lives.
This protocol details the preparation of senescent cells in parallel with similarly growth-arrested quiescent cells in culture, followed by metabolic labeling with pSILAC. Cells are then harvested, homogenized into lysates, and processed for mass spectrometry acquisition and analysis. The data obtained from mass spectrometry are then used to determine protein half-lives using a simplified quantitative method employing a single time point and half-life calculations performed in a spreadsheet. Using this approach, estimates of protein half-lives can be measured in a comprehensive and quantitative manner that is more authentic to unperturbed cellular conditions than protocols that use blockers of protein synthesis or turnover.

<table><tbody><tr><td>Acetonitrile (LC-MS grade)</td><td>Grainger</td><td>AH015</td><td></td></tr><tr><td>Ammonium Bicarbonate</td><td>Millipore-Sigma</td><td>9830</td><td></td></tr><tr><td>Antibiotic-Antimycotic (100x)</td><td>ThermoFisher</td><td>15240062</td><td></td></tr><tr><td>BCA Assay Kit</td><td>ThermoFisher</td><td>23227</td><td></td></tr><tr><td>Dithiothreotol (DTT)</td><td>Sigma</td><td>D9779</td><td></td></tr><tr><td>DMEM, high glucose, HEPES</td><td>ThermoFisher</td><td>12430112</td><td></td></tr><tr><td>dNTP Mix</td><td>ThermoFisher</td><td>R0191</td><td></td></tr><tr><td>Fetal Bovine Serum, certified, heat inactivated</td><td>ThermoFisher</td><td>10082147</td><td></td></tr><tr><td>Formic Acid</td><td>Sigma</td><td>27001</td><td></td></tr><tr><td>Gammacel 40 Exactor</td><td>Best Theratronics</td><td></td><td>Cesium Irradiator for cells</td></tr><tr><td>GlycoBlue</td><td>ThermoFisher</td><td>AM2238</td><td></td></tr><tr><td>Iodoacetamide (IAA)</td><td>Sigma</td><td>I1149</td><td>Light sensitive</td></tr><tr><td>IMR-90 primary lung fibroblasts</td><td>ATCC</td><td>CCL-186</td><td></td></tr><tr><td>iRT Kit (indexed retention time)</td><td>Biognosys</td><td>Ki-3002-2</td><td>Indexed Retention Time Peptide Standards</td></tr><tr><td>Isopropanol</td><td>ThermoFisher</td><td>423835000</td><td></td></tr><tr><td>Mascot</td><td>Matrix Science</td><td>Mascot Daemon 2.8</td><td>Proteomic database searching software</td></tr><tr><td>Maxima Reverse Transcritase (200 U/&amp;micro;L)</td><td>ThermoFisher</td><td>EP0742</td><td></td></tr><tr><td>MEM Non-Essential Amino Acids Solution (100x)</td><td>ThermoFisher</td><td>11140050</td><td></td></tr><tr><td>Nano LC System</td><td>ThermoFisher</td><td>ULTIM3000RSLCNANO</td><td></td></tr><tr><td>Oasis HLB Solid Phase Extraction Cartirdges</td><td>Waters</td><td>186000383</td><td></td></tr><tr><td>Orbitrap Mass Spectrometer</td><td>ThermoFisher</td><td>Q Exactive HF Orbitrap</td><td></td></tr><tr><td>Phenol/Chloroform/Isoamyl alcohol (25:24:1), 100 mM EDTA, pH 8.0</td><td>ThermoFisher</td><td>327110025</td><td></td></tr><tr><td>Phosphate Buffered Saline (PBS)</td><td>ThermoFisher</td><td>10010023</td><td></td></tr><tr><td>Pierce SILAC Protein Quantitation Kit (Trypsin) -DMEM</td><td>ThermoFisher</td><td>A33972</td><td></td></tr><tr><td>QuantStudio 6 Real-Time PCR System</td><td>ThermoFisher</td><td></td><td></td></tr><tr><td>Random Hexamer Primer</td><td>ThermoFisher</td><td>SO142</td><td></td></tr><tr><td>Senescence &amp;beta;-Galactosidase Staining Kit</td><td>Cell Signaling</td><td>9860</td><td></td></tr><tr><td>Skyline</td><td>University of Washington</td><td>Skyline-Daily v21.2.1.424</td><td>Free and open source qantiative proteomic software. Available on www.skyline.ms</td></tr><tr><td>Sonicator waterbath</td><td>Branson</td><td>CPX-952-516R</td><td></td></tr><tr><td>TRIzol Reagent</td><td>ThermoFisher</td><td>15596018</td><td>Referred to as phenol in text; hazardous</td></tr><tr><td>TRYPle Express</td><td>ThermoFisher</td><td>12605010</td><td></td></tr><tr><td>Trypsin (sequencing grade)</td><td>Promega</td><td>V5113</td><td></td></tr><tr><td>TURBO Dnase (2U/ uL)</td><td>ThermoFisher</td><td>AM2238</td><td></td></tr><tr><td>Urea</td><td>ThermoFisher</td><td>29700</td><td></td></tr><tr><td>Water (LC-MS grade)</td><td>Grainger</td><td>AH365</td><td></td></tr></tbody></table>

measurement of protein turnover rates in senescent and non-dividing cultured cells with metabolic labeling and mass spectrometry

Mounting evidence has shown that the accumulation of senescent cells in the central nervous system contributes to neurodegenerative disorders such as Alzheimer&#39;s and Parkinson&#39;s diseases. Cellular senescence is a state of permanent cell cycle arrest that typically occurs in response to exposure to sub-lethal stresses. However, like other non-dividing cells, senescent cells remain metabolically active and carry out many functions that require unique transcriptional and translational demands and widespread changes in the intracellular and secreted proteomes. Understanding how protein synthesis and decay rates change during senescence can illuminate the underlying mechanisms of cellular senescence and find potential therapeutic avenues for diseases exacerbated by senescent cells. This paper describes a method for proteome-scale evaluation of protein half-lives in non-dividing cells using pulsed stable isotope labeling by amino acids in cell culture (pSILAC) in combination with mass spectrometry. pSILAC involves metabolic labeling of cells with stable heavy isotope-containing versions of amino acids. Coupled with modern mass spectrometry approaches, pSILAC enables the measurement of protein turnover of hundreds or thousands of proteins in complex mixtures. After metabolic labeling, the turnover dynamics of proteins can be determined based on the relative enrichment of heavy isotopes in peptides detected by mass spectrometry. In this protocol, a workflow is described for the generation of senescent fibroblast cultures and similarly arrested quiescent fibroblasts, as well as a simplified, single-time point pSILAC labeling time-course that maximizes coverage of anticipated protein turnover rates. Further, a pipeline is presented for the analysis of pSILAC mass spectrometry data and user-friendly calculation of protein degradation rates using spreadsheets. The application of this protocol can be extended beyond senescent cells to any non-dividing cultured cells such as neurons.

This protocol describes a method to globally compare protein half-lives between senescent and non-dividing, quiescent control cells using pSILAC and minimal time points. This protocol details generation of senescent and quiescent cells in culture, metabolic labeling of cells with stable isotopes of arginine and lysine over 3 days, quantifying the relative abundances of heavy and light peptide isotopes by mass spectrometry, and a straightforward and accessible calculation of protein half-lives using spreadsheet formulas (Figure 1). This method is highly flexible and can be adapted to numerous cell types and conditions.
As part of the generation of senescent cells for this protocol, two methods of senescence validation are used: SA-&#946;Gal-positive cells visualized by microscopy and increased levels of senescent markers quantified using RT-qPCR analysis. The measurement of senescent markers should yield clear distinctions between quiescent and senescent cells for comparisons of the two cells populations to be considered valid. For SA-&#946;Gal activity, senescent cells should appear blue while quiescent control cells have no or very little color (Figure 2A). This assay can be quantified by counting the positive, blue-stained cells as a percentage of the total number of cells, and then comparing the percentage positivity rate between quiescent control and senescent cells. It is important that this protocol be performed at the same time for the comparison of both cell states; SA-&#946;Gal activity relies on the pH of the staining solution, so the results can vary substantially between assays and should be considered a qualitative measure.
The RT-qPCR analysis of senescent markers will show high levels of the mRNAs encoding SASP factors (IL6, CXCL8, IL1B) and the cell cycle inhibitors (CDKN2A/p16, CDKN1A/p21) in most senescent models. Although some variation is expected based on cell type, culture condition, and senescent inducer41,42, most senescent cells display greater than five-fold higher levels of IL6, CXCL8, and CDKN1A/p21 mRNAs compared to quiescent control cells; conversely, the levels of LMNB1 and PCNA mRNAs, encoding proliferation markers, should be low or absent in senescent cells compared with quiescent&#160;cells (Figure 2B). Taken together, a significant percentage of SA-&#946;Gal positivity and the expected expression of a panel of senescence-associated mRNA markers are sufficient to confirm the induction of senescence in an experiment.
Processing protein samples for mass spectrometry analysis consists of in-solution digestion (2 days) and solid-phase extraction (4-6 h). To perform untargeted proteomic analysis, the resulting peptides are submitted for LC-MS/MS analysis using data-dependent acquisition (DDA). On Orbitrap instruments, a DDA method can be specified in the mass spectrometry instrument software method editor. The mass spectrometer settings used in this study are shown in Figure 3A. Liquid chromatography settings can also be specified within the method editor. For this study, a 90 min linear gradient with increasing organic phase (acetonitrile) was used (Figure 3B). Following a successful acquisition, the total ion current (TIC) should contain an intense signal during the linear gradient portion of the method (Figure 3C). This protocol describes an example protocol on an Orbitrap instrument, but the calculation of protein half-lives can be performed on data obtained from any mass spectrometer that was collected in DDA-mode, which is available on several types of instruments (e.g., Orbitraps and time-of-flight instruments) and vendors. The acquisition settings will be unique to the type and configuration of the instrument used, and it is recommended to use the settings suggested by the mass spectrometry facility. This method is also compatible with non-DDA methods (SRM, PRM, DIA), as long as quantitative chromatographic peak areas for heavy and light peaks can be extracted from the raw data files and entered into the spreadsheet formulas.
Raw mass spectrometry files are searched with one of the many available proteomic database search tools to identify peptides and proteins. For example, this study utilized the Mascot43 search engine. To obtain chromatographic peak areas for quantification of heavy and light peptides, database search results are imported into a proteomic software capable of chromatographic peak areas such as Skyline39,40. Examination of the extracted ion chromatograms of peptides (Figure 4) will reveal the relative proportion of heavy and light peptide signals. A lower proportion of heavy peptide signal relative to light peptide signal in senescent cells indicates a slower protein turnover rate (Figure 4A), and a higher heavy peptide signal relative to light indicates a faster protein turnover rate (Figure 4B). The unlabeled samples should show little or no heavy peptide signal. Any apparent heavy peptide signal in the unlabeled samples is considered background noise and will be subtracted during the final calculations. Quantification of chromatographic peak areas for light and heavy peptides for all treatment and labeling conditions must be exported for subsequent calculation of protein turnover rates and statistical analysis.
By using single time point analysis, the quantification of protein half-lives is simple to perform and can be done conveniently in spreadsheets. In Table 3, the half-lives for 695 proteins identified from the mass spectrometry analysis were calculated. Starting from protein-level heavy and light isotope abundances for each protein in the samples (input for the 1) Raw Data sheet), a percent heavy isotope (termed the Ratio, R) is then automatically calculated on the sheet titled 2) Ratio H | H + L. At this step, the R from heavy labeled samples is normalized by subtracting the R from unlabeled samples to produce a final R for the quiescent and senescent triplicates. Since unlabeled samples should not have any exogenously added heavy isotope, they are used to eliminate the background signal. From R, the kdeg (turnover rate) is calculated using the following formula:
<img alt="Equation 1" src="/files/ftp_upload/63835/63835eq01.jpg" style="margin: auto;" />
The half-life (H, in days) is determined for each protein in the quiescent control and senescent triplicates (sheet titled 3) Half-Life (days)) using the following formula:
<img alt="Equation 2" src="/files/ftp_upload/63835/63835eq02.jpg" style="margin: auto;" />
These half-lives are then averaged among the quiescent and senescent triplicates to generate a quiescent and senescent mean half-life for each protein as well as a p-value. Calculated half-lives are then filtered for values that are negative, which occurs when the heavy signal from light-labeled cells (the background) is greater than the heavy signal from heavy-labeled cells. This filtering resulted in 707 proteins from 841 identified with valid half-lives for the results presented here.
The half-life is then reported as a log2 ratio of senescent over quiescent control cells (log2FC) and plotted in a volcano plot (Figure 5). For example, by looking at the 5) Analysis sheet, the coagulation II thrombin receptor (F2R) protein can be seen to have a half-life in quiescent cells of 0.51 days (column B) and a half-life in senescent cells of 1.07 days (column C) that yielded a log2FC of ~1.06 (column D) with a p-value of 0.001.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63835/63835fig01.jpg" /> 
Figure 1: Diagram of pSILAC workflow for senescent and quiescent control cells. Human IMR-90 fibroblasts were used to prepare quiescent (low-serum) or senescent (IR) cell cultures for comparison of protein half-lives. Cells were then labeled with SILAC Light or SILAC Heavy media with isotopic arginine and lysine for 3 days. Lysates were extracted from cells, digested, desalted, and analyzed using mass spectrometry. Heavy and light peptide isotope peaks correspond to newly synthesized and pre-existing peptides, respectively. Half-lives were calculated using an equation for exponential decay. <a href="https://www.jove.com/files/ftp_upload/63835/63835fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63835/63835fig02.jpg" /> 
Figure 2: Validation of senescent phenotypes using SA-&#946;Gal and RT-qPCR analyses. (A) Senescent and quiescent cells are re-plated at the time of harvest into a 6-well plate and stained for SA-&#946;Gal using the SA-&#946;Gal staining kit. Blue color in the cell body is positive for senescence. Images are taken in brightfield with color at 10x magnification, size marker in red. (B) RT-qPCR analysis comparing senescent cells (red) and cycling cells (gray) from an unrelated experiment. Increases in the levels of CDKN1A/p21, CXCL8, and IL6 mRNAs are indicative of senescence, as is the decrease in LMNB1 mRNA levels. <a href="https://www.jove.com/files/ftp_upload/63835/63835fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63835/63835fig03.jpg" /> 
Figure 3: Representative methods for data-dependent acquisition (DDA) scans and liquid chromatography gradient of pSILAC cultures on the Q-Exactive HF orbitrap mass spectrometer. (A) Recommended instrument settings in the instrument software for data-dependent analysis of whole-cell lysates from a pSILAC experiment. (B) Example liquid chromatography flow gradient method settings. Peptides are eluted over a 90-min linear gradient ranging from 5% to 35% buffer B (0.2 % formic acid and 99.8% acetonitrile), followed by a 10 min wash with 80% buffer B, and 25 min of equilibration with 5% buffer B. (C) A representative total ion chromatogram (TIC) of a mass spectrometry acquisition of IMR-90 fibroblast peptides acquired with the specified settings. <a href="https://www.jove.com/files/ftp_upload/63835/63835fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63835/63835fig04.jpg" /> 
Figure 4: Representative extracted ion chromatograms of peptides with altered turnover during senescence. (A) Chromatographic peak areas of the peptide FQMTQEVVCDECPNVK++ from the protein DnaJ homolog subfamily B member 11 (DNAJB11) in senescent and non-senescent cells. Following 3 days of SILAC, senescent cells incorporate less heavy isotope into this peptide compared with quiescent (non-senescent) cells, as indicated by a reduction in the peak area of heavy isotope-containing peptide (blue) relative to the light peptide (red), indicating that this peptide has reduced turnover in senescent cells. (B) Chromatographic peak areas of the peptide VQAQVIQETIVPK++ from the protein Splicing Factor 3a Subunit 1 (SF3A1) in senescent and non-senescent cells. Following 3 days of SILAC, senescent cells incorporate a higher proportion of heavy isotope into this peptide compared with quiescent (non-senescent) cells, as indicated by a reduction in the peak area of heavy isotope-containing peptide (blue) relative to the light peptide (red), indicating that this peptide has increased turnover in senescent cells. The unlabeled (Day 0) conditions show no incorporation of heavy isotope for both peptides, as expected. <a href="https://www.jove.com/files/ftp_upload/63835/63835fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/63835/63835fig05.jpg" /> 
Figure 5: Comparison of protein half-lives in senescent and quiescent cells determined from pSILAC labeling. (A) Volcano plot displaying the log2 ratio of senescent / control (quiescent) for each of the 695 identified proteins; in this experiment, Light and Heavy labeling media contained no glucose and no phenol red. (B) Tables showing the top 10 proteins with the most increased or decreased half-lives in quiescent cells versus senescent cells (left and right, respectively). <a href="https://www.jove.com/files/ftp_upload/63835/63835fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Table 1: Media and buffers used in this protocol. <a href="https://www.jove.com/files/ftp_upload/63835/Table 1-63835R1.xlsx" target="_blank">Please click here to download this Table.</a>
Table 2: RT-qPCR primers used in this protocol. <a href="https://www.jove.com/files/ftp_upload/63835/Table 2-63835R1.xlsx" target="_blank">Please click here to download this Table.</a>
Table 3: SILAC analysis workbook for calculation of protein half-lives, fold changes, and t-tests. <a href="https://www.jove.com/files/ftp_upload/63835/Table 3 v3.xlsx" target="_blank">Please click here to download this Table.</a>

Watch this Scientific Journal Video about Measurement of Protein Turnover Rates in Senescent and Non-Dividing Cultured Cells with Metabolic Labeling and Mass Spectrometry at JoVE.com

Measurement of Protein Turnover Rates in Senescent and Non-Dividing Cultured Cells with Metabolic Labeling and Mass Spectrometry

This protocol describes the workflow for metabolic labeling of senescent and non-dividing cells with pulsed SILAC, untargeted mass spectrometry analysis, and a streamlined calculation of protein half-lives.

Mounting evidence has shown that the accumulation of senescent cells in the central nervous system contributes to neurodegenerative disorders such as ...

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Biochemistry

3.4K Views.  National Institutes of Health. Hoe cellen eiwithomeostase behouden, is niet goed begrepen in de context van cellulaire stress en senescentie. In dit protocol bieden we een eenvoudige en nauwkeurige manier om de eiwitomzet te berekenen, wat een belangrijk onderdeel is van eiwithomeostase in levende cellen. Pulse SILAC geeft een experimentator de mogelijkheid om de eiwitomzet voor individuele eiwitten en het gehele proteoom te meten op een high-throughput manier die ook authentieker is voor biologische systemen dan andere me...

Video: Measurement of Protein Turnover Rates in Senescent and Non-Dividing Cultured Cells with Metabolic Labeling and Mass Spectrometry

Mass Spectrometry and Luminogenic-based Approaches to Characterize Phase I Metabolic Competency of In Vitro Cell Cultures

Xenobiotic metabolizing enzymes play a key function in the biotransformation of medicines and toxicants by adding functional groups that increase solubility and facilitate excretion. On some occasions those structural modifications lead to the formation of new toxic products. In order to reduce animal testing, chemical risk can be assessed using metabolically competent cells. The expression of metabolic enzymes, however, is not stable over time in many in vitro primary culture systems and is often partial or absent in cell lines. Therefore, the study of medicines, additives, and environmental pollutants metabolism in vitro should ideally be conducted in cell systems where metabolic activity has been characterized. We explain here an approach to measure the activity of a class of metabolic enzymes (Human Phase I) in 2D cell lines and primary 3D cultures using chemical probes and their metabolic products quantifiable by UPLC mass spectrometry and luminometry. The method can be implemented to test the metabolic activity in cell lines and primary cells derived from a variety of tissues.

Mass Spectrometry and Luminogenic-based Approaches to Characterize Phase I Metabolic Competency of In Vitro Cell Cultures

Metabolic competency of in vitro systems is a key requirement for the biotransformation and disposition of drugs and toxicants. In this protocol, we describe the application of reference metabolic probes to assess phase I metabolism in cell cultures.

Xenobiotic metabolizing enzymes play a key function in the biotransformation of medicines and toxicants by adding functional groups that increase ...

Single-Cell Quantification of Protein Degradation Rates by Time-Lapse Fluorescence Microscopy in Adherent Cell Culture

Proteins are in a dynamic state of synthesis and degradation and their half-lives can be adjusted under various circumstances. However, most commonly used approaches to determine protein half-life are either limited to population averages from lysed cells or require the use of protein synthesis inhibitors. This protocol describes a method to measure protein half-lives in single living adherent cells, using SNAP-tag fusion proteins in combination with fluorescence time-lapse microscopy. Any protein of interest fused to a SNAP-tag can be covalently bound by a fluorescent, cell permeable dye that is coupled to a benzylguanine derivative, and the decay of the labeled protein population can be monitored after washout of the residual dye. Subsequent cell tracking and quantification of the integrated fluorescence intensity over time results in an exponential decay curve for each tracked cell, allowing for determining protein degradation rates in single cells by curve fitting. This method provides an estimate for the heterogeneity of half-lives in a population of cultured cells, which cannot easily be assessed by other methods. The approach presented here is applicable to any type of cultured adherent cells expressing a protein of interest fused to a SNAP-tag. Here we use mouse embryonic stem (ES) cells grown on E-cadherin-coated cell culture plates to illustrate how single cell degradation rates of proteins with a broad range of half-lives can be determined.

This protocol describes a method to determine protein half-lives in single living adherent cells, using pulse labeling and fluorescence time-lapse imaging of SNAP-tag fusion proteins.

Proteins are in a dynamic state of synthesis and degradation and their half-lives can be adjusted under various circumstances. However, most commonly used ...

Developmental Biology

A Sensitive Method to Quantify Senescent Cancer Cells

Human cells do not indefinitely proliferate. Upon external and/or intrinsic cues, cells might die or enter a stable cell cycle arrest called senescence. Several cellular mechanisms, such as telomere shortening and abnormal expression of mitogenic oncogenes, have been shown to cause senescence. Senescence is not restricted to normal cells; cancer cells have also been reported to senesce.
	Chemotherapeutical drugs have been shown to induce senescence in cancer cells. However, it remains controversial whether senescence prevents or promotes tumorigenesis. As it might eventually be patient-specific, a rapid and sensitive method to assess senescence in cancer cell will soon be required.
	To this end, the standard &beta;-galactosidase assay, the currently used method, presents major drawbacks: it is time consuming and not sensitive. We propose here a flow cytometry-based assay to study senescence on live cells. This assay offers the advantage of being rapid, sensitive, and can be coupled to the immunolabeling of various cellular markers.

Whether senescence prevents or promotes tumorigenesis remains controversial. Since chemotherapeutical drugs can induce cancer cells to senesce, studying senescence is essential for proposing new therapies. However, the standard and broadly used &beta;-galactosidase assay presents major drawbacks. We propose here a rapid and sensitive flow cytometry-based assay to quantify senescence.

Human cells do not indefinitely proliferate. Upon external and/or intrinsic  cues, cells might die or enter a stable cell cycle arrest called senescence. ...

Medicine

Induction and Validation of Cellular Senescence in Primary Human Cells

Cellular senescence is a state of permanent cell cycle arrest activated in response to different damaging stimuli. Activation of cellular senescence is a hallmark of various pathophysiological conditions including tumor suppression, tissue remodeling and aging. The inducers of cellular senescence in vivo are still poorly characterized. However, a number of stimuli can be used to promote cellular senescence ex vivo. Among them, most common senescence-inducers are replicative exhaustion, ionizing and non-ionizing radiation, genotoxic drugs, oxidative stress, and demethylating and acetylating agents. Here, we will provide detailed instructions on how to use these stimuli to induce fibroblasts into senescence. This protocol can easily be adapted for different types of primary cells and cell lines, including cancer cells. We also describe different methods for the validation of senescence induction. In particular, we focus on measuring the activity of the lysosomal enzyme Senescence-Associated &#946;-galactosidase (SA-&#946;-gal), the rate of DNA synthesis using 5-ethynyl-2&#39;-deoxyuridine (EdU) incorporation assay, the levels of expression of the cell cycle inhibitors p16 and p21, and the expression and secretion of members of the Senescence-Associated Secretory Phenotype (SASP). Finally, we provide example results and discuss further applications of these protocols.

Here, we discuss a series of protocols for induction and validation of cellular senescence in cultured cells. We focus on different senescence-inducing stimuli and describe the quantification of common senescence-associated markers. We provide technical details using fibroblasts as a model, but the protocols can be adapted to various cellular models.

Cellular senescence is a state of permanent cell cycle arrest activated in response to different damaging stimuli. Activation of cellular senescence is a ...

Metabolic Labeling and Membrane Fractionation for Comparative Proteomic Analysis of Arabidopsis thaliana Suspension Cell Cultures

Plasma membrane microdomains are features based on the physical properties of the lipid and sterol environment and have particular roles in signaling processes. Extracting sterol-enriched membrane microdomains from plant cells for proteomic analysis is a difficult task mainly due to multiple preparation steps and sources for contaminations from other cellular compartments. The plasma membrane constitutes only about 5-20% of all the membranes in a plant cell, and therefore isolation of highly purified plasma membrane fraction is challenging. A frequently used method involves aqueous two-phase partitioning in polyethylene glycol and dextran, which yields plasma membrane vesicles with a purity of 95% 1. Sterol-rich membrane microdomains within the plasma membrane are insoluble upon treatment with cold nonionic detergents at alkaline pH. This detergent-resistant membrane fraction can be separated from the bulk plasma membrane by ultracentrifugation in a sucrose gradient 2. Subsequently, proteins can be extracted from the low density band of the sucrose gradient by methanol/chloroform precipitation. Extracted protein will then be trypsin digested, desalted and finally analyzed by LC-MS/MS. Our extraction protocol for sterol-rich microdomains is optimized for the preparation of clean detergent-resistant membrane fractions from Arabidopsis thaliana cell cultures.
We use full metabolic labeling of Arabidopsis thaliana suspension cell cultures with K15NO3 as the only nitrogen source for quantitative comparative proteomic studies following biological treatment of interest 3. By mixing equal ratios of labeled and unlabeled cell cultures for joint protein extraction the influence of preparation steps on final quantitative result is kept at a minimum. Also loss of material during extraction will affect both control and treatment samples in the same way, and therefore the ratio of light and heave peptide will remain constant. In the proposed method either labeled or unlabeled cell culture undergoes a biological treatment, while the other serves as control 4.

Metabolic Labeling and Membrane Fractionation for Comparative Proteomic Analysis of Arabidopsis thaliana Suspension Cell Cultures

Here we describe a robust method for the fractionation of plant plasma membranes into detergent resistant and detergent soluble membranes based on a mixture of unlabeled and in vivo fully 15N labeled Arabidopsis thaliana cell cultures. The procedure is applied for comparative proteomic studies to understand signaling processes.

Plasma membrane microdomains are features based on the physical properties of the lipid and sterol environment and have particular roles in signaling ...

Biology

Selected Reaction Monitoring Mass Spectrometry for Absolute Protein Quantification

Absolute quantification of target proteins within complex biological samples is critical to a wide range of research and clinical applications. This protocol provides step-by-step instructions for the development and application of quantitative assays using selected reaction monitoring (SRM) mass spectrometry (MS). First, likely quantotypic target peptides are identified based on numerous criteria. This includes identifying proteotypic peptides, avoiding sites of posttranslational modification, and analyzing the uniqueness of the target peptide to the target protein. Next, crude external peptide standards are synthesized and used to develop SRM assays, and the resulting assays are used to perform qualitative analyses of the biological samples. Finally, purified, quantified, heavy isotope labeled internal peptide standards are prepared and used to perform isotope dilution series SRM assays. Analysis of all of the resulting MS data is presented. This protocol was used to accurately assay the absolute abundance of proteins of the chemotaxis signaling pathway within RAW 264.7 cells (a mouse monocyte/macrophage cell line). The quantification of Gi2 (a heterotrimeric G-protein &#945;-subunit) is described in detail.

This protocol describes how to perform absolute quantification assays of target proteins within complex biological samples using selected reaction monitoring. It was used to accurately quantify proteins of the mouse macrophage chemotaxis signaling pathway. Target peptide selection, assay development, and qualitative and quantitative assays are described in detail.

Absolute quantification of target proteins within complex biological samples is critical to a wide range of research and clinical applications. This ...

Techniques to Induce and Quantify Cellular Senescence

In response to cellular stress or damage, proliferating cells can induce a specific program that initiates a state of long-term cell-cycle arrest, termed cellular senescence. Accumulation of senescent cells occurs with organismal aging and through continual culturing in vitro. Senescent cells influence many biological processes, including embryonic development, tissue repair and regeneration, tumor suppression, and aging. Hallmarks of senescent cells include, but are not limited to, increased senescence-associated &#946;-galactosidase activity (SA-&#946;-gal); p16INK4A, p53, and p21 levels; higher levels of DNA damage, including &#947;-H2AX; the formation of Senescence-associated Heterochromatin Foci (SAHF); and the acquisition of a Senescence-associated Secretory Phenotype (SASP), a phenomenon characterized by the secretion of a number of pro-inflammatory cytokines and signaling molecules. Here, we describe protocols for both replicative and DNA damage-induced senescence in cultured cells. In addition, we highlight techniques to monitor the senescent phenotype using several senescence-associated markers, including SA-&#946;-gal, &#947;-H2AX and SAHF staining, and to quantify protein and mRNA levels of cell cycle regulators and SASP factors. These methods can be applied to the assessment of senescence in various models and tissues.

Cellular senescence, the irreversible state of cell-cycle arrest, can be induced by various cellular stresses. Here, we describe protocols to induce cellular senescence and methods to assess markers of senescence.

In response to cellular stress or damage, proliferating cells can induce a specific program that initiates a state of long-term cell-cycle arrest, termed ...

Measuring Diurnal Rhythms in Autophagic and Proteasomal Flux

Cells employ several methods for recycling unwanted proteins and other material, including lysosomal and non-lysosomal pathways. The main lysosome-dependent pathway is called autophagy, while the primary non-lysosomal method for protein catabolism is the ubiquitin-proteasome system. Recent studies in model organisms suggest that the activity of both autophagy and the ubiquitin-proteasome system is not constant across the day but instead varies according to a daily (circadian) rhythm. The ability to measure biological rhythms in protein turnover is important for understanding how cellular quality control is achieved and for understanding the dynamics of specific proteins of interest. Here we present a standardized protocol for quantifying autophagic and proteasomal flux in vivo that captures the circadian component of protein turnover. Our protocol includes details for mouse handling, tissue processing, fractionation, and autophagic flux quantification using mouse liver as the starting material.

We describe our protocol for measuring biological rhythms in protein catabolism via autophagy and the proteasome in mouse liver.

Cells employ several methods for recycling unwanted proteins and other material, including lysosomal and non-lysosomal pathways. The main ...

In Vivo Quantification of Protein Turnover in Aging C. Elegans using Photoconvertible Dendra2

Proteins are synthesized and degraded constantly within a cell to maintain homeostasis. Being able to monitor the degradation of a protein of interest is key to understanding not only its life cycle, but also to uncover imbalances in the proteostasis network. This method shows how to track the degradation of the disease-causing protein huntingtin. Two versions of huntingtin fused to Dendra2 are expressed in the C. elegans nervous system: a physiological version or one with an expanded and pathogenic stretch of glutamines. Dendra2 is a photoconvertible fluorescent protein; upon a short ultraviolet (UV) irradiation pulse, Dendra2 switches its excitation/emission spectra from green to red. Similar to a pulse-chase experiment, the turnover of the converted red-Dendra2 can be monitored and quantified, regardless of the interference from newly synthesized green-Dendra2. Using confocal-based microscopy and due to the optical transparency of C. elegans, it is possible to monitor and quantify the degradation of huntingtin-Dendra2 in a living, aging organism. Neuronal huntingtin-Dendra2 is partially degraded soon after conversion and cleared further over time. The systems controlling degradation are deficient in the presence of mutant huntingtin and are further impaired with aging. Neuronal subtypes within the same nervous system exhibit different turnover capacities for huntingtin-Dendra2. Overall, monitoring any protein of interest fused to Dendra2 can provide important information not only on its degradation and the players of the proteostasis network involved, but also on its location, trafficking, and transport.

In Vivo Quantification of Protein Turnover in Aging C. Elegans using Photoconvertible Dendra2

Presented here is a protocol to monitor degradation of the protein huntingtin fused to the photoconvertible fluorophore Dendra2.

Proteins are synthesized and degraded constantly within a cell to maintain homeostasis. Being able to monitor the degradation of a protein of interest is ...

Metabolic Labeling

Metabolic labeling is used to probe the biochemical transformations and modifications that occur in a cell. This is accomplished by using chemical analogs that mimic the structure of natural biomolecules. Cells utilize analogs in their endogenous biochemical processes, producing compounds that are labeled. The label allows for the incorporation of detection and affinity tags, which can then be used to elucidate metabolic pathways using other biochemical analytical techniques, such as SDS-PAGE and NMR.
This video introduces the concepts of metabolic labeling and show two general procedures.&#160; The first uses isotopic-labeling, to characterize the phosphorylation of a protein. The second covers a photoreactive labeling to characterize protein-protein interaction within a Also three applications of metabolic labeling are presented: labeling plant material, labeling RNA to measure kinetics and labeling glycans in developing embryos.

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Metabolic labeling is used to investigate the machinery of a cell. This is accomplished using chemical analogs to probe the biochemical transformations and modifications that occur. This video will show the principles of metabolic labeling, typical isotopic and photoreactive labeling procedures, and some applications.
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Metabolic labeling can be conducted using a number of strategies. Here we will describe isotopic, photoreactive, and bio-orthogonal labeling.
Isotopic labeling is performed using structural analogs that are chemically identical to their natural counterparts, but have uncommon isotopes incorporated into their structure. In this L-lysine analog the carbon and nitrogen atoms are replaced with carbon-13 and nitrogen-15. Cells cultured in the presence of isotopic analogs will incorporate them into their biochemical structures. Metabolites are collected from the cells and purified for analysis. Samples with stable isotopes are analyzed using techniques such as mass spectrometry or NMR spectroscopy. Samples with radioactive labels are analyzed using liquid scintillation counting and x-ray films, which will be demonstrated in the isotopic labeling protocol.
Photoreactive labels are functional groups incorporated into proteins, which are stable until exposed to ultraviolet light. The functional group forms a reactive radical, which binds to the nearest protein. A common example, L-photo-leucine, contains a diazirine ring, which is a photoreactive crosslinker. In contrast to isotopic labeling, there is some chemical dissimilarity between photo-reactive chemical analogs and their natural counterparts. Cells may preferentially incorporate natural compounds over analogs. Therefore, it is important to perform photo-reactive labeling in medium free of the compound being mimicked. Once exposed to ultraviolet radiation, the photo-reactive groups in a labeled protein become unstable and highly reactive, causing it to cross-link with interacting proteins, creating a protein complex. Cross-linked complexes, act as snapshots that can then be analyzed using SDS-PAGE and mass spectrometry methods. This provides insights into what reactions are occurring in the metabolic pathway by identifying reaction species and how they interact by determining binding sites.
Bioorthogonal labeling strategies utilize analogs with small functional groups that have little to no reactivity with natural biomolecules. For example, azides are small functional groups, whose reactivity is said to be orthogonal to biochemical reactions. In the Staudinger ligation, a phosphine group attacks the azido group. This yields a transition state that intramolecularly reacts with a nearby ester, resulting in an amine-bonded ligand. Bioorthogonal functional groups incorporated into biomolecules can be ligated with detection tags such as fluorescent functional groups, and affinity tags such as antigens.
Now that some concepts and strategies for metabolic labeling have been discussed, let&#39;s look at the process in the laboratory.
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The first step in a metabolic labeling experiment is to collect the protein of interest. To do this, cells are grown on a plate, and an expression method is used to promote synthesis of the desired protein. In this example, leucine-rich repeat kinases, or LRRK, are expressed. Disodium phosphate, containing radioactive phosphorus-32, is used as the analog. Proper measures must be taken to protect against ionizing radiation. This includes setting up a work area, wearing proper protective equipment, and checking for radioactive contamination. Once safety measures have been taken, the medium containing the isotopic analogs is prepared. The medium from the culture is removed and, replaced with one containing the isotopic chemical analogs and then incubated. Following incubation, the cells are lysed. The lysate is collected and purified.
After purification, proteins are resolved using SDS-PAGE and then transferred to a PVDF membrane. Autoradiography is performed by exposing the membrane to x-ray film and measured using a phosphor imager. Western blotting is used to measure relative protein levels in the PVDF membrane. In this example the phosphorylation levels of leucine-rich repeat kinases synthesized in 293T cells were measured. The autoradiogram shows how much phosphorous was incorporated into the protein. Western blotting elucidates the levels of the LRRK proteins. Image analysis software is used to obtain quantitative data of phosphorylation levels of the proteins.
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In this next procedure, photoreactive labeling is demonstrated. First, the cells are prepared and cultured. The photoreactive analog is added to the cells at the mid-log phase and incubated. In this procedure p-benzoylphenylalanine is used. Samples are collected over intervals and put on ice. The samples are then exposed to obtain snapshots of the biochemical pathways over time. The proteins of interest are then purified and resolved using SDS-PAGE.
A photoreactive labeling strategy was used to identify compounds that interact with the protein of interest. Immunodetection with Western blotting shows protein bands that indicate higher molecular weight proteins are present in the irradiated samples. These are from cross-linking due to protein-protein interaction occurring during the UV irradiation.
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Now that we&#39;ve reviewed metabolic labeling procedures, let&#39;s look at some of the ways the process is used.
Metabolic labeling concepts can be extended to multicellular organisms. Plants are grown in a sealed environment, rich in stable isotopes to produced labeled plant material. Carbon dioxide containing carbon-13 is added to the enclosure, while nitrogen-15 rich fertilizer is used. The resulting harvested plant material can help answer questions about carbon and nitrogen cycling from the ecosystem.
Labeling enables the separation of newly synthesized RNA from older RNA. By changing the initial concentration of analog, the kinetics of new RNA synthesis can be determined. The results show that the concentration of 4-thiouridine affects how much new RNA is transcribed. Additionally, incorporation rates of the label into RNA can be directly quantified with a spectrophotometer.
Using biorthogonal click chemistry, glycans in a zebra fish embryo can be labeled. The eggs are injected with a labeling compound that results in alkyne labels on the glycans. The glycans in the larvae are then ligated to a dye compound at the desired development stage. The glycans in the embryos are then imaged. Glycans produced at different time points can be identified by labeling using different colors at different stages of embryo development.
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You&#39;ve just watched JoVE&#39;s video on metabolic labeling. This video described the concepts behind metabolic labeling and their strategies, went over two general procedures, and covered some of the uses of the techniques.
Thanks for watching!
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Isotopic, Photoreactive and Bioorthogonal Metabolic Labeling

Metabolic labeling is used to probe the biochemical transformations and modifications that occur in a cell. This is accomplished by using chemical analogs ...

Measurement of Protein Turnover Rates in Senescent and Non-Dividing Cultured Cells with Metabolic Labeling and Mass Spectrometry

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