From the Laboratory of Proteomics and Analytical Technologies, SAIC-Frederick, Inc., National Cancer Institute, Frederick, MD 21702;
Expression Pathology Incorporated, Gaithersburg, MD 20877; ¶ Armed Forces Institute of Pathology, Department of Genitourinary Pathology, Washington, DC 20306; and || Naval Research Laboratory, Washington, DC 20375
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ABSTRACT |
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Tissue-based proteomic studies are inherently attractive for relating protein biomarkers directly to disease. Whereas fresh and/or frozen tissue samples may represent attractive samples from which proteomic biomarker investigations may be conducted, they are often difficult to obtain in large numbers and relatively expensive to store in a stable form. Formalin fixation and paraffin embedding of tissue is the standard processing methodology practiced in pathology laboratories worldwide, resulting in a highly stable form of tissue that is easily stored due to its inherent stability at room temperature. Hence, formalin-fixed paraffin-embedded (FFPE)1 tissues represent a potentially attractive resource for conducting retrospective protein biomarker investigation. Arising from the high degree of covalently cross-linked proteins in FFPE tissue, immunohistochemistry (IHC) is currently the only published technology capable of providing "proteomic" information from these samples, and unfortunately IHC methods lack sensitivity, quantitation, and scalability. Methodologies designed to increase IHC sensitivity by decreasing the effects of formalin-induced protein cross-linking have shown some success, yet quantitation and scalability issues remain (6, 7). In addition, IHC requires a priori knowledge of individual proteins being analyzed, thereby limiting biomarker discovery approaches. Other methods to procure and assay soluble proteins directly from FFPE tissue have also shown limited success on a small scale (810). Development of the capability to conduct large scale analyses of tissues using proteomics approaches analogous to the scale and throughput of high throughput gene expression analysis could have far reaching implications on protein biomarker investigations of disease through interrogation of the vast archived FFPE tissue collections.
Laser-based tissue microdissection has enabled molecular analysis of specifically defined populations of cells captured directly from their tissue microenvironment (3, 1113). Despite recent interest in conducting tissue microdissection of FFPE tissues for mRNA extraction and analysis, microdissection of FFPE tissues is not widely practiced due to lack of technological optimization for fixed tissue and lack of methodologies for extraction and analysis of soluble protein (1417).
Dramatic improvements in MS instrumentation and the rapid growth of genomic databases have enabled development of high throughput proteomic approaches to identify and quantify large numbers of proteins from complex samples such as cancer cell lysates (5, 1821). Identification of proteins by MS in cells obtained by microdissection of fresh frozen tissue has recently been demonstrated (22, 23). The combined application of tissue microdissection and MS analysis of FFPE tissue has the potential for generating protein biomarker data necessary for dissecting those elements key to the pathological manifestation of cancer and other diseases.
We have utilized a novel tissue microdissection technique optimized for the extraction of cells from FFPE tissue and a biochemical methodology for the recovery of soluble proteins and peptides suitable for analysis by nanoflow reversed-phase liquid chromatography (nanoRPLC)-MS/MS for a proteomic investigation of benign prostate hyperplasia (BPH) and prostate cancer (PCa) tissue. The data presented demonstrate the potential for identification of proteins and their relative expression levels and relate this information to a particular disease state from FFPE tissue. This study represents the first differential large scale proteome analysis from FFPE-archived tissue. Furthermore, this study provides a technological framework by which the vast archive of pathologically characterized FFPE clinical samples may be analyzed utilizing state of the art MS-based proteomic technologies.
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MATERIALS AND METHODS |
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Tissue Pathology and Processing
A FFPE whole-mount prostate case was chosen and histologically analyzed by standard methods. 5-µm-thick sections were cut and placed on glass slides and stained with hematoxylin and eosin for identification of histologically distinct tissue regions. Histological analysis was performed on an Olympus BX51 microscope, and images (10 x 20 magnification) were taken with an Olympus Q color camera mounted on the microscope. The IHC analysis for prostate-specific antigen (PSA) was performed using standard methodologies with a mouse monoclonal anti-PSA primary antibody (Dako Cytomation, Carpinteria, CA) and standard colorimetric detection methods.
For tissue microdissection, 10-µm-thick tissue sections were cut from the FFPE whole-mount prostate tissue block, placed on the coated slides, and heated for 60 min at 58 °C. Paraffin was removed by treatment in SubX organic solvent (Surgipath Medical Industries, Richmond, IL) twice for 5 min; this was followed by tissue rehydration through multiple, graded ethanol solutions and distilled water. Tissue was counterstained with Mayers hematoxylin, dehydrated through graded ethanol solutions, and air-dried. Tissue was rehydrated prior to microdissection with 50% glycerol in water for 5 min. Slides were placed upside down below the 10x objective and visualized to locate cellular regions with specific histological features, which were mapped using the accompanying stage software. The microdissection was automated, allowing for modulation of the laser and scanning of the receiving substrate via computer-aided design/computer-aided machining (CAD/CAM). Microdissection was performed by software directed laser pulses to strike at a constant velocity and rate throughput over the previously defined and mapped cellular regions to achieve complete transfer of cells within the selected area into a 1.5-ml low binding microcentrifuge receiving tube.
The tissue microdissection instrument is based on an optical setup designed to direct a pulsed laser onto a target slide holding a 10-µm-thick tissue section. An excimer laser (MPB Technologies PSX-100) operating at the following conditions was utilized for microdissection: 248 nm wavelength, 2.5 ns pulse, Emax = 5 mJ, repetition rate = 0.1100 Hz. Target slides are made of optically transparent quartz with an energy transfer coating with the exact dimensions of a standard histology glass slide. The slide stage is a computer controlled, XY translation stage with a maximum translation speed of 200 mm/s. The laser is split by a 1/8 beam splitter to an energy meter, and the remaining beam travels to an UV reflective mirror and directed down (Z) to a 10x microscope objective (LMU-10x-UVR, OFR), which focuses the laser onto the slide and allows for observation of the dissection process via a confocally aligned CCD camera.
To assess the impact of formalin fixation on the ability to identify peptides extracted from FFPE tissue sections, a liver was procured from a freshly sacrificed male C57 black mouse. A lobe of the liver was cut into two equal sections. One section of the liver was frozen on dry ice and placed at 80 °C prior to embedding in optimal cutting temperature compound (OCT) and cryosectioned. The other section was immediately placed in 10% buffered formalin for 96 h prior to embedding in paraffin and sectioning. The frozen tissue sections were placed in 70% ethanol for 30 s and then distilled water for 30 s to remove the OCT, followed by dehydration in 70, 85, and 100% ethanol and quickly air dried. The FFPE liver tissue was deparaffinized in two changes of SubX for 5 min followed by rehydration through graded ethanol solutions and distilled water. The entire sections from both the formalin-fixed and frozen tissues were placed separately in 20 µl of LT-MS buffer, processed, and analyzed by nanoRPLC-MS/MS as described below.
Protein Extraction
Cells collected by microdissection for array analysis and nanoRPLC-MS/MS analysis were processed by reagents according to the manufacturers recommendations (Expression Pathology, Inc.). Cellular material for array analysis was suspended in 20 µl of LT-ID buffer A, incubated at 95 °C for 90 min, then cooled on ice for 3 min at which time 1 µl of LT-ID buffer B was added followed by incubation at 37 °C for 1 h. Samples were stored at 20 °C until analysis. Cellular material for nanoRPLC-MS/MS analysis was suspended in 20 µl of LT-MS reaction buffer, incubated at 95 °C for 90 min, then cooled on ice for 3 min at which time 1 µl of trypsin (1518 U) was added followed by incubation at 37 °C overnight. DTT was added to a final concentration of 10 mM, and the samples were heated for 5 min at 95 °C to reduce cysteine residues. Digestates were stored at 20 °C until analysis.
Protein Microarray Assay
Two-fold serial dilutions of protein extracts were arrayed in duplicate on a nitrocellulose membrane using a manual arrayer (V&P Scientific, Inc., San Diego, CA). A protein dot blot assay for PSA expression was performed using standard immunoassay conditions with a mouse monoclonal anti-PSA primary antibody (DakoCytomation) and a biotinylated anti-mouse IgG secondary antibody (Pierce Biotechnology, Inc.), streptavidin-alkaline phosphatase (Pierce Biotechnology, Inc.), and nitroblue tetrazolium/5-bromo-4-chloro-3-indolyl phosphate substrate (Moss, Inc., Pasadena, MD) for colorimetric signal detection. Images of the membrane were captured by a CCD camera as a TIFF file and analyzed with image analysis software (GridGrinder, Corning Incorporated, Corning, NY) for spot intensity. The data were imported into a Microsoft Excel for graphing and statistical analysis
Sample Preparation and Trypsin-mediated 18O Labeling
Tryptic peptides generated from BPH and PCa cells were desalted using C-18 ZipTip microcolumns (Millipore, Billerica, MA), lyophilized to dryness, and resuspended in 0.1% TFA for nanoRPLC-MS/MS analysis. For the isotope labeling, protein extracts from an approximately equivalent number of microdissected cells were lyophilized and reconstituted separately in H216O (BPH) and H218O (PCa) each containing 20% methanol (v/v). Porcine sequencing grade trypsin (Promega) was resuspended in the appropriately labeled water and added to each sample at an enzyme-to-protein ratio of 1:20. Samples were incubated for 16 h at 37 °C, after which an additional equivalent aliquot of the trypsin solution was added, and the samples were incubated for an additional 6 h. Because tryptic peptides are produced via the extraction of the proteins from the cells dissected from the FFPE tissues, the role of trypsin during this step is to enzymatically exchange the C-terminal carboxyl oxygen atoms with the appropriate isotope. TFA was added to a final concentration of 0.4% (v/v), and the solutions were boiled for 10 min. Samples were pooled, lyophilized to dryness, and resuspended in 0.1% TFA prior to analysis.
NanoRPLC-MS/MS Analysis
NanoRPLC was performed using an Agilent 1100 capillary LC system (Agilent Technologies, Palo Alto, CA) coupled on-line to either a linear ion trap (LIT) mass spectrometer (LTQ, Thermo Electron, San Jose, CA) or a hybrid LIT-FT-ICR MS. NanoRPLC separations of each sample were performed using 75-µm inner diameter x 360 outer diameter x 10-cm-long fused silica capillary columns (Polymicro Technologies, Phoenix, AZ) that were slurry packed in-house with 3 µm, 300 Å pore size C-18 silica bonded stationary phase (Vydac, Hysperia, CA). After injecting 1 µl of sample, the column was washed for 30 min with 98% mobile-phase A (0.1% formic acid in water) at a flow rate of 0.5 µl/min. Peptides were eluted using a linear gradient of 2% mobile-phase B (0.1% formic acid in ACN) to 40% solvent B in 110 min, then to 98% B in an additional 30 min, all at a constant flow rate of 0.25 µl/min.
The LIT-MS was operated in a data-dependent MS/MS mode in which each full MS scan is followed by five MS/MS scans where the five most abundant peptide molecular ions are selected for CID using a normalized collision energy of 35%. Dynamic exclusion was utilized to minimize redundant selection of peptides previously selected for CID. The heated capillary temperature and electrospray voltage were set at 160 °C and 1.5 kV, respectively. Data were collected over a broad mass to charge (m/z) precursor ion selection scan range of 4002000 followed by eight segmented precursor selection scan ranges (e.g. gas phase fractionation in the m/z dimension, GPFm/z) using the following overlapping m/z intervals: m/z 400605, 595805, 7951005, 9951205, 11951405, 13951605, 15951805, 17952000, 400805, and 7951200. Data for the 16O/18O-labeled BPH/PCa experiment was acquired using FT-ICR detection in centroid mode for the full MS scan (m/z 2002000) at 100,000 resolution followed by MS/MS using the LIT on the top five molecular ions detected in the FT-ICR scan.
Bioinformatic Analysis
Tandem mass spectra were searched against the UniProt human and mouse proteomic databases (01/04/05 release) from the European Bioinformatics Institute (www.ebi.ac.uk/integr8) using SEQUEST (Thermo Electron). Peptides were searched using fully tryptic cleavage constraints and a dynamic 4.008-amu modification on the C terminus for the 18O isotope-labeling analysis. For a peptide to be considered legitimately identified, it had to achieve stringent charge state and proteolytic cleavage-dependent cross-correlation (Xcorr) scores of 1.9 for [M+H]1+, 2.2 for [M+2H]2+, and 3.5 for [M+3H]3+ (3.1 for the 18O-labeling analysis), and a minimum delta correlation
score (Cn) of 0.08. SEQUEST results were further filtered using software developed in-house to determine unique peptides and proteins. To evaluate false positive identifications, the entire PCa dataset was searched against a reversed human protein database in a similar fashion as previously described (24). The calculations reveal that using the SEQUEST parameters described above results in less than 1.5% of the identifications in each dataset being classified as a false positive.
The molecular function and cellular localization of the unique proteins identified in the various analyses performed in this study were categorized by gene ontology (www.geneontology.org). GRAVY values for each unique protein were calculated according to the method of Kyte and Doolittle (25).
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RESULTS |
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Tissue Microdissection
10-µm-thick prostate tissue sections were cut and placed on single energy transfer-coated slides for microdissection. Approximately 200,000 cells from each of the two PSA+ tumor regions (PCa 1 and 2), 200,000 cells from each of the two PSA+ BPH regions (BPH 1 and 2), and 100,000 cells from the PSAstromal region were microdissected (Fig. 1C) using a novel tissue microdissection technique developed specifically for FFPE tissue, termed the ExCelleratorTM. The energy transfer layer between the slide and tissue section imparts a photoabsorption process where each incident laser pulse removes a three-dimensional pixel of tissue from the quartz slide, propelling it into a receiving tube. The amount of tissue transferred in a single pixel is a function of the diameter of the laser, the tissue thickness, and the laser fluence. The software controlling the moveable stage that held the tissue slide was programmed to perform CAD/CAM microdissection of specific demarcated cellular regions for microdissection. Laser repetition rates were at a constant 99 Hz, allowing for transfer of all cells in a marked region within seconds. A CCD camera positioned in line with the optical setup allowed for tissue visualization, identification of cell populations for microdissecting, and real-time analysis/monitoring of the transfer process. The total number of microdissected cells was approximated by determining the area of the lesion marked for CAD/CAM-controlled microdissection, the approximate diameter of a single cancer cell (15 µm), and thickness of the section (10 µm).
Protein Microarray Analysis
Protein extracts were prepared for microarray analysis from the tumor epithelial PCa 1 and stromal regions, serially diluted, and spotted on a microarray in duplicate along with buffer. Approximately 90 cell equivalents of protein were spotted at the high end of the 2-fold dilution series. Standard immunoassay detection of the protein array with a monoclonal anti-PSA antibody indicates positive detection of PSA in the sample prepared from the PSA+ epithelial cancer area (PCa 1), whereas no PSA was detected in the sample prepared from the PSA stromal area (results shown graphically in Fig. 1D). This indicates the ability to detect PSA expression in a microarray format and demonstrates the dilutability of extracted protein from formalin-fixed tissue. A duplicate parallel array assayed without primary PSA antibody showed no signal (data not shown). All samples were normalized for protein concentration.
Analysis of Peptides by MS
To investigate the potential utility of conventional MS-based proteomic methods for conducting identification of peptides extracted from FFPE PCa tissue, 200,000 cells from PCa 2 and BPH 2 regions (Fig. 1, A and C) were microdissected as described and analyzed by nanoRPLC-MS/MS. Approximately 100,000 cell equivalents of protein from each extract were analyzed by nanoRPLC-MS/MS, resulting in the identification of over 1,300 and 2,200 unique, fully tryptic peptides representing 702 and 1,156 unique proteins, from the BPH and PCa FFPE tissue extracts, respectively (supplemental Tables 1 and 2). Of these proteins, roughly 25% were identified by two or more unique peptides in both tissue extracts (Fig. 2, A and B).
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To determine if formalin fixation impacts trypsin enzymatic specificity, the numbers of peptides identified from the PCa tissue analysis with arginyl and lysyl termini were tabulated. The results show that the percentage of peptides identified with arginyl or lysyl termini is 56 and 44%, respectively. Considering that the prevalence of these two residues within proteins is roughly equivalent (i.e. 5.7% of all residues) the most probable outcome would have been a 50:50 distribution of peptides identified containing arginyl and lysyl termini. Although the present results do not exactly match the theoretically predicted distribution, they do not differ significantly enough to draw a reasonable conclusion, suggesting that formalin fixation does not have a negative impact on the enzyme specificity of trypsin.
The number of identified peptides that contain internal missed tryptic cleavage sites was also evaluated. Approximately 25% of the identified peptides contain an internal missed cleavage site. Although this percentage is at the high end, this result may be caused by the fact that certain regions of the proteins may not be completely unfolded or denatured, and therefore some arginyl and lysyl residues are not fully accessible to trypsin.
The molecular functions and cellular localization of the proteins identified in the analysis of the FFPE PCa and BPH tissue samples were classified using gene ontology as shown in Fig. 4. The identified proteins were found to have a broad range of molecular functions and arose from every cell compartment. The gene ontology classification of the proteins identified in these two samples was compared with the proteins within the entire human protein database. This analysis demonstrates that there is no significant bias in the spectrum of proteins identifiable from FFPE tissues. Whereas the human proteome database is predicted to contain a higher percentage of membrane proteins than observed in the present result from the proteomic analysis of FFPE PCa and BPH tissue, this observation is not surprising considering the present sample preparation methodology lacks any specific protocol for membrane protein enrichment. In the case of signal transduction proteins, the analysis of the FFPE tissues revealed a lower percentage than that classified within the human proteome. It is important to point out, however, that signal transduction proteins are considered to be low abundant proteins, and therefore it would be expected that they would be somewhat underrepresented in this analysis compared with their preponderance in the human proteome. The range of hydropathy values for the proteins identified from the PCa and BPH tissues was also calculated (25). The proteins identified in each set show similar results and covered the complete range of GRAVY values (i.e. 2.5 to +2.5). A majority of the proteins are slightly hydrophilic in nature (calculated hydropathy values of 0 to 1), slightly lower than that observed for the entire human protein database (data not shown). In general, these results are consistent with other global proteomic analyses performed in our laboratory (26, 27).2 A comparison of the data shows that a global proteome analysis of FFPE tissue samples utilizing the described methodology results in a broad coverage of protein localization and function.
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DISCUSSION |
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Analysis of the BPH-microdissected tissue extract yielded similar results with a few notable distinctions. A number of the prostatic markers identified in the cancer extract were identified by similar numbers of unique peptides in the BPH extract. Indeed, PSA was identified by an additional unique peptide in BPH, and IHC revealed the presence of PSA in both cancer and BPH regions from the same tissue section. This is not surprising because PSA has been under significant scrutiny for its inability to accurately distinguish between the two prostatic diseases. This is further supported by the quantitative proteomic analysis utilizing 18O isotope labeling where PSA peptides were identified with approximately equivalent expression levels in each tissue based on the observed 16O/18O isotope ratio (Fig. 5).
In the case of MIC-1, while four peptides were identified in the PCa-microdissected tissue extracts, no peptides were identified in the BPH-microdissected tissue extract, suggesting it is more abundant in PCa cells than BPH. Indeed, lack of MIC-1 protein expression has been associated with BPH (39), whereas high levels of MIC-1 expression have been linked to early prostate carcinogenesis (40). This result is further corroborated by the quantitative proteomic analysis where MIC-1 expression was detected in the cancer extract by isotopic labeling, whereas no MIC-1 expression was observed in the BPH extract, suggesting absence of expression in this case of BPH. In addition, a further 68 proteins were identified as differentially expressed between PCa and BPH by isotopic-labeling analysis.
FFPE tissues are a largely unexplored archive in MS-based proteomics. Therefore, it is important to understand the effects of formalin fixation and long-term storage on the samples. Unfortunately it is extremely difficult to identify unpredictable modifications within a complex mixture. Our results, however, show that just over 6% of the identified peptides contained a formylated lysine residue, and the percentage of oxidized methionine residues or peptides containing missed tryptic cleavages is not significantly different than that observed in proteomic studies of fresh cells (24, 26). Because we do not presently understand all of the possible modifications that may be observed in proteins extracted from FFPE tissues, biological information will primarily be obtained from the analysis of unmodified peptides.
The ability to conduct proteomic investigations of FFPE tissue has the potential to provide invaluable information regarding disease states and may aid in the development of improved clinical characterization and/or diagnostic techniques. Historically, the study of FFPE tissues has been limited to analysis of protein and RNA expression through use of affinity reagents such as antibodies or RNA probes (IHC and in situ hybridization). The methodology described here enables large scale proteomic technologies, such as MS, to be applied to vast archives of clinical and research tissues that had been previously thought of as intractable to such analyses. Use of tissue microdissection provides the ability to accurately correlate protein expression to histological information for further investigation of a variety of proteins and their involvement in disease. In contrast to existing tissue microdissection approaches, the novel microdissection technology described here has been developed and optimized for procurement of cells directly from standard formalin-fixed tissue sections for downstream protein analysis. In addition, this technology is a noncontact method that explodes cells directly downward into a collection tube providing the added benefit of CAD/CAM automation for high throughput tissue microdissection capabilities.
The accurate diagnosis of prostate cancer is extremely challenging using current methodologies, and it is likely to benefit from using a panel of biomarkers. Indeed, the assessment of MIC-1 protein expression in conjunction with PSA testing may allow for distinction between BPH and frank PCa, alleviating incorrect diagnoses and further unnecessary tests.
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FOOTNOTES |
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1 The abbreviations used are: FFPE, formalin-fixed paraffin-embedded; IHC, immunohistochemistry; LT, Liquid TissueTM; nanoRPLC, nanoflow reversed-phase liquid chromatography; BPH, prostate hyperplasia; CCD, charge-coupled device; PCa, prostate cancer; PSA, prostate-specific antigen; CAD/CAM, computer-aided design/computer-aided machining; OCT, optimal cutting temperature compound; LIT, linear ion trap.
2 B. L. Hood, M. M. Darfler, T. G. Guiel, B. Furusato, D. A. Lucas, B. R. Ringeisen, I. A. Sesterhenn, T. P. Conrads, T. D. Veenstra, and D. B. Krizman, unpublished data.
* This work was supported by federal funds from the National Cancer Institute, National Institutes of Health, under contract NO1-CO-12400. The Costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
S The on-line version of this article (available at http://www.mcponline.org) contains supplemental material.
Published, MCP Papers in Press, August 9, 2005, DOI 0.1074/mcp.M500102-MCP200.
** To whom correspondence may be addressed: Laboratory of Proteomics and Analytical Technologies, National Cancer Institute, SAIC-Frederick, Inc., P.O. Box B, Frederick, MD 21702. Tel.: 301-846-7286; Fax: 301-846-6037; E-mail: veenstra{at}ncifcrf.gov
To whom correspondence may be addressed: Expression Pathology Inc., 9290 Gaither Rd., Gaithersburg, MD 20877. Tel.: 301-977-3654; Fax: 301-926-9283; E-mail: d.krizman{at}expressionpathology.com
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REFERENCES |
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