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Obi, E.N.; Tellock, D.A.; Thomas, G.J.; Veenstra, T.D. Protein Extraction Methods from Formalin-Fixed Paraffin-Embedded Tissue. Encyclopedia. Available online: https://encyclopedia.pub/entry/45585 (accessed on 15 June 2024).
Obi EN, Tellock DA, Thomas GJ, Veenstra TD. Protein Extraction Methods from Formalin-Fixed Paraffin-Embedded Tissue. Encyclopedia. Available at: https://encyclopedia.pub/entry/45585. Accessed June 15, 2024.
Obi, Ekenedirichukwu N., Daniel A. Tellock, Gabriel J. Thomas, Timothy D. Veenstra. "Protein Extraction Methods from Formalin-Fixed Paraffin-Embedded Tissue" Encyclopedia, https://encyclopedia.pub/entry/45585 (accessed June 15, 2024).
Obi, E.N., Tellock, D.A., Thomas, G.J., & Veenstra, T.D. (2023, June 14). Protein Extraction Methods from Formalin-Fixed Paraffin-Embedded Tissue. In Encyclopedia. https://encyclopedia.pub/entry/45585
Obi, Ekenedirichukwu N., et al. "Protein Extraction Methods from Formalin-Fixed Paraffin-Embedded Tissue." Encyclopedia. Web. 14 June, 2023.
Protein Extraction Methods from Formalin-Fixed Paraffin-Embedded Tissue
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The relatively developments in mass spectrometry (MS) have provided novel opportunities for this technology to impact modern medicine. One of those opportunities is in biomarker discovery and diagnostics. Key developments in sample preparation have enabled a greater range of clinical samples to be characterized at a deeper level using MS. While most of these developments have focused on blood, tissues have also been an important resource. Fresh tissues, however, are difficult to obtain for research purposes and require significant resources for long-term storage. There are millions of archived formalin-fixed paraffin-embedded (FFPE) tissues within pathology departments worldwide representing every possible tissue type including tumors that are rare or very small. Owing to the chemical technique used to preserve FFPE tissues, they were considered intractable to many newer proteomics techniques and primarily only useful for immunohistochemistry. In the past couple of decades, however, researchers have been able to develop methods to extract proteins from FFPE tissues in a form making them analyzable using state-of-the-art technologies such as MS and protein arrays. 

formalin-fixed paraffin-embedded tissues mass spectrometry biomarker discovery disease diagnosis

1. Introduction

Over the past two decades the number of proteins identified in global proteomic studies using liquid chromatography combined with mass spectrometry (LC-MS) has increased from a few hundred to several thousand [1][2]. Even the identification of proteins within serum and plasma, which is notoriously difficult to characterize owing to its wide range of protein concentrations, increased from about 500 to over 4000 proteins [3][4]. This increase catalyzed the rapid increase in the number of studies focused on identifying biomarkers within blood [5]. While the number of studies exponentially increased, the number of clinically validated biomarkers did not. The reasons for this are both technical and physiological. Blood samples are incredibly complex containing proteins that originate from virtually every area of the body through being actively secreted or leaking from dying or diseased cells [6]. While these processes provide blood with a rich diversity of proteins, this diversity is dominated by 22 proteins that make up about 99% of its protein content [3]. This domination makes detecting the lowest abundant proteins, which are anticipated to contain the highest yield of biomarkers, technically challenging owing to the limited dynamic range of analytical instrumentation [7]. While methods such as high abundant protein depletion, chromatographic separation, data-independent analysis, etc. have enabled greater coverage of lower abundant proteins, they cannot overcome the physiological barriers in finding disease-specific biomarkers in blood [8][9][10].
The physiological barriers to identifying biomarkers arise from the ubiquitous nature of the interaction between blood and the body. No cell is more than four cell units removed from the circulatory system and cellular proteins are constantly being dumped into the bloodstream. While a biomarker’s concentration may be significantly elevated at the site from which it is excreted, its concentration will be diluted at the point where the sample is removed from the patient. This dilution may eliminate any quantitative difference in the biomarker’s level between healthy and disease-affected patients. Another barrier is recognition of the biomarker’s origin. While studies analyzing both blood and tissue from patients to correlate biomarker measurements in both types of samples [11] may help, they do not absolutely prove the source of the biomarker.

2. Protein Extraction Methods 

In retrospect, it is somewhat puzzling why it took so many years for formalin-fixed paraffin-embedded (FFPE) tissues to become a commonly used sample in MS-based proteomic analysis. It was related to two reasons: (1) many proteomic laboratories were unaware of the vast archive of tissue that was available and (2) it was thought that the formalin-fixation made it impossible to retrieve analyzable protein. In the end, most MS-based proteomic studies used extraction methods based on antigen retrieval (AR) methods originally developed for IHC, which simply use a combination of buffer and heat [12]. To enhance the recovery of proteins for MS analysis, researchers have used various combinations of heat and buffers that contain detergents and/or reducing agents such as dithiothreitol or glycine [13][14][15][16]. As the popularity of FFPE tissue in proteomic analysis grew several companies including Qiagen, Hilder, Germany, Agilent Technologies, Santa Clara, CA, USA, Invent Biotechnologies, Inc., Plymouth, MN, USA, Covaris, Woburn, MA, USA and Bio Basic, Markham, ON, USA develop kits that could be used to help simplify and standardize protein extraction.
One concern when using FFPE tissues is to what extent are formalin crosslinks reversed during sample preparation. This question is generally answered through the direct comparison of matching FFPE and fresh frozen tissues. As described above, most sample preparation methods use a combination of buffer, detergent, reducing agents, and heat, with some also using elevated pressure. Studies comparing matched FFPE and fresh frozen tissue have generally shown that these conditions are sufficient to reverse a majority of protein crosslinks. For example, a study by Addis et al. [17] extracted proteins from FFPE and fresh frozen sheep tissues by immersing tissue sections in buffer comprised of 20 mM Tris HCl, 2% SDS, and 200 mM dithiothreitol (DTT) (pH 8.8) and heating the sample at 100 °C for 20 min, followed by additional heating at 80 °C for 2 h. Comparison of the samples using SDS-PAGE, Western blotting, and LC-MS showed similar results suggesting that the reversal of the crosslinks does not impact the ability to identify peptides obtained from the intact proteins. Gene ontology analysis of the identified proteins showed no significant bias in the proteins identified in the FFPE or fresh frozen tissues.
Another study extracted proteins from matching FFPE and fresh frozen mouse livers using 0.1 M Tris HCL, 10 mM sodium deoxycholate, and 10 mM sodium lauroyl sarcosinate (pH 9.0) and heated the samples at 95 °C for 60 min [18]. The samples were also subjected to 60 cycles at 45,000 psi for 95 s, followed by 5 s at atmospheric pressure. Finally, the samples were subjected to 50 cycles at 45,000 psi for 20 s and 15 s at atmospheric pressure. This research showed that the addition of pressure cycling technology (PCT) to the sample preparation more than tripled the amount of protein extracted from the FFPE tissue, increasing the extraction efficiency to almost 100%.
So how do these combinations of buffers, detergents, heat, and pressure reverse formalin crosslinks? The key is hydration. The prevailing hypothesis is that heating the FFPE tissue in the presence of a buffer and detergent denatures that protein molecules and allows water molecules to access cavities that will hydrolyze the formalin-protein bond [19]. This hypothesis is consistent with the observation that heat treatment combined with high pressure increases the extent of the reversal of formalin-protein crosslinks by increasing the level of hydration within the protein’s interior [20][21][22].
In the early days of developing methods for extracting proteins from FFPE tissues for proteomic analysis, Fowler et al. (2007) performed a series of studies to evaluate the use of different buffers, detergents, reducing agents, and temperatures on the extraction efficiency [23]. In these studies, they prepared a FFPE surrogate tissue using a known concentration of lysozyme, ribonuclease A or a 1:2 molar ratio of carbonic anhydrase and lysozyme. The study showed that heat, a denaturant, and a detergent were all necessary for optimal protein extraction. Sodium dodecyl sulfate (SDS) was the single most important ingredient for protein extraction. A buffer containing 2% SDS had a protein extraction efficiency 13-fold greater than buffers without SDS. While Triton X-100 is an often-used protein detergent, it was not very effective on extracting FFPE proteins. Heating the tissues at either 80 °C for 2 h or 100 °C for 20 min was the most effective for extracting proteins. The addition of reducing agents such as glycine showed a modest benefit in increasing protein extraction. The optimal pH range was found to be between 4–6, however, results using model proteins of various isoelectric points (pI) suggested that extraction of any individual protein was dependent on the pH and pI of the specific protein.
This same group was also instrumental in showing the value of pressure in extracting proteins from FFPE tissues [24][25]. Using a FFPE lysozyme sample, they extracted the protein at pressures ranging from 14.7 to 50,000 psi. The results showed that extraction of the FFPE sample using a buffer and SDS at 40,000 psi increased the percentage of protein extracted from the lysozyme sample from 96% from only 26% observed when the extraction was performed at atmospheric pressure (i.e., 14.7 psi). The effect of using higher pressure for extraction on the LC-MS profiles (extracted ion chromatograms and protein identifications) was also evaluated [25]. Proteins were extracted from a multi-protein FFPE tissue surrogates in Tris-HCl buffer with 2% (w/v) SDS at both 40,000 and 14.7 psi, digested with trypsin, and analyzed using LC/MS. At 14.7 psi and pH4, only lysozyme and RNase A were identified using MS/MS, while none of the component proteins were correctly identified at pH 8. Increasing the extraction pressure to 40,000 psi resulted in the identification of all five surrogate proteins at both pH 4 and 8, with sequence coverages ranging from 28% to 69%. These results were comparable to those obtained when the surrogate protein mixture was analyzed prior to fixation. Besides enhanced peptide identification, the false identification rates for the pressure extracted samples were only 5.7% (pH 8) and 7.8% (pH 4), compared to the rates for the non-pressure extracted tissue surrogates of 42% (pH 4) and 100% (pH 8). The MS spectra of the native protein mixture, pressure-extracted, and non-pressure extracted multi-protein surrogate samples showed differences in protein quality. The unfixed protein mixture spectrum (panel A) showed several well resolved peaks eluting as did the profile for the tissue surrogate extracted under elevated pressure (panel B). Many of the peaks within the spectrum of the non-pressure treated FFPE surrogate mixture (panel C) were reduced in intensity and eluted later, which suggests that much of the protein material remained cross-linked and was not completely digested.

References

  1. Shen, Y.; Jacobs, J.M.; Camp, D.G.; Fang, R.; Moore, R.J.; Smith, R.D.; Xiao, W.; Davis, R.W.; Tompkins, R.G. Ultra-high-efficiency strong cation exchange LC/RPLC/MS/MS for high dynamic range characterization of the human plasma proteome. Anal. Chem. 2002, 76, 1134–1144.
  2. Müller, J.B.; Geyer, P.E.; Colaço, A.R.; Treit, P.V.; Strauss, M.T.; Oroshi, M.; Doll, S.; Virreira Winter, S.; Bader, J.M.; Kohler, N.; et al. The proteome landscape of the kingdoms of life. Nature 2020, 582, 592–596.
  3. Tirumalai, R.S.; Chan, K.C.; Prieto, D.A.; Issaq, H.J.; Conrads, T.P.; Veenstra, T.D. Characterization of the low molecular weight human serum proteome. Mol. Cell. Proteom. 2003, 2, 1096–1103.
  4. Uyar, D.S.; Huang, Y.W.; Chesnik, M.A.; Doan, N.B.; Mirza, S.P. Comprehensive serum proteomic analysis in early endometrial cancer. J. Proteom. 2021, 234, 104099–104107.
  5. Mendes, M.L.; Dittmar, G. Targeted proteomics on its way to discovery. Proteomics 2022, 22, e2100330.
  6. Anderson, N.L.; Anderson, N.G. The human plasma proteome: History, character, and diagnostic prospects. Mol. Cell. Proteom. 2002, 1, 845–867.
  7. Lee, P.Y.; Osman, J.; Low, T.Y.; Jamal, R. Plasma/serum proteomics: Depletion strategies for reducing high-abundance proteins for biomarker discovery. Bioanalysis 2019, 11, 1799–1812.
  8. Krasny, L.; Huang, P.H. Data-independent acquisition mass spectrometry (DIA-MS) for proteomic applications in oncology. Mol. Omics 2021, 17, 29–42.
  9. Makawita, S.; Diamandis, E.P. The bottleneck in the cancer biomarker pipeline and protein quantification through mass spectrometry-approaches: Current strategies for candidate verification. Clin. Chem. 2010, 56, 212–222.
  10. Kulyyassov, A.; Fresnais, M.; Longuespee, R. Targeted liquid chromatography-tandem mass spectrometry analysis of proteins: Basic principles, applications, and perspectives. Proteomics 2021, 21, e2100153.
  11. Johann Jr, D.J.; Wei, B.H.; Prieto, D.A.; Chan, K.C.; Ye, X.; Valera, V.A.; Simpson, R.M.; Rudnick, P.A.; Xiao, Z.; Issaq, H.J.; et al. Combined blood/tisse analysis for cancer biomarker discovery: Application to renal cell carcinoma. Anal. Chem. 2010, 82, 1584–1588.
  12. Shi, S.R.; Cote, R.J.; Taylor, C.R. Antigen retrieval techniques: Current perspectives. J. Histochem. Cytochem. 2001, 49, 931–937.
  13. Mantsiou, A.; Makridakis, M.; Fasoulakis, K.; Katafigiotis, I.; Constantinides, C.A.; Zoidakis, J.; Roubelakis, M.G.; Vlahou, A.; Lygirou, V. Proteomics analysis of formalin fixed paraffin embedded tissues in the investigation of prostate cancer. J. Proteome Res. 2020, 19, 2631–2642.
  14. Azimzadeh, O.; Barjaktarovic, Z.; Aubele, M.; Calzada-Wack, J.; Sarioglu, H.; Atkinson, M.J.; Tapio, S. Formalin-fixed paraffin-embedd (FFPE) proteome analysis using gel-free and gel-based proteomics. J. Proteome Res. 2010, 9, 4710–4720.
  15. Sun, R.; Hunter, C.; Chen, C.; Ge, W.; Morrice, N.; Liang, S.; Zhu, T.; Yuan, C.; Ruan, G.; Zhang, Q.; et al. Accelerated protein biomarker discovery from FFPE tissue samples using single-shot, short gradient microflow SWATH-MS. J. Proteome Res. 2020, 19, 2732–2741.
  16. Fu, Z.; Yan, K.; Rosenberg, A.; Jin, Z.; Crain, B.; Athas, G.; Heide, R.S.; Howard, T.; Everett, A.D.; Herrington, D.; et al. Improved protein extraction and protein identification from archival formalin-fixed paraffin-embedded human aortas. Proteom. Clin. Appl. 2013, 7, 217–224.
  17. Addis, M.F.; Tanca, A.; Pagnozzi, D.; Crobu, S.; Fanciulli, G.; Cossu-Rocca, P.; Uzzau, S. Generation of high-quality protein extracts from formalin-fixed, paraffin-embedded tissues. Proteomics 2009, 9, 3815–3823.
  18. Uchida, Y.; Sasaki, H.; Terasaki, T. Establishet ad validation of highly accurate formalin-fixed paraffin-embedded quantitative proteomics by heat-compatible pressure cycling technology using phase-transfer surfactant and SWATH-MS. Sci. Rep. 2020, 10, 11271.
  19. Fowler, C.B.; O’Leary, T.J.; Mason, J.T. Improving the proteomic analysis of archival tissue by using pressure-assisted protein extraction: A mechanistic approach. J. Proteom. Bioinform. 2014, 7, 151–157.
  20. Refaee, M.; Tezuka, T.; Akasaka, K.; Williamson, M.P. Pressure-dependent changes in the solution structure of hen egg-white lysozyme. J. Mol. Biol. 2003, 327, 857–865.
  21. Frye, K.J.; Royer, C.A. Probing the contribution of internal cavities to the volume change of protein unfolding under pressure. Protein Sci. 1998, 7, 2217–2222.
  22. Kobashigawa, Y.; Sakurai, M.; Nitta, K. Effect of hydrostatic pressure on unfolding of alphalactalbumin: Volumetric equivalence of the molten globule and unfolded state. Protein Sci. 1999, 8, 2765–2772.
  23. Fowler, C.B.; Cunningham, R.E.; O’Leary, T.J.; Mason, J.T. ‘Tissue surrogates’ as a model for archival formalin-fixed paraffin-embedded tissues. Lab. Investig. 2007, 87, 836–846.
  24. Fowler, C.B.; Cunningham, R.E.; Waybright, T.J.; Blonder, J.; Veenstra, T.D.; O’Leary, T.J.; Mason, J.T. Elevated hydrostatic pressure promotes protein recovery from formalin-fixed, paraffin-embedded tissue surrogates. Lab. Investig. 2008, 88, 185–195.
  25. Fowler, C.B.; Chesnick, I.E.; Moore, C.D.; O’Leary, T.J.; Mason, J.T. Elevated pressure improves the extraction and identification of proteins recovered from formalin-fixed paraffin-embedded tissue surrogates. PLoS ONE 2010, 5, e14253.
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